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Copyright ©1983 Paul Stamets and J.S. Chilton. All rights reserved. No
part of this book may be reproduced or transmitted in any form by any
means without written permission from the publisher, except by a reviewer,
who may quote brief passages in a review.

Produced by Paul Stamets and J.S. Chilton

Published by Agarikon Press

Box 2233, Olympia, Washington, 98507

Western Distribution by Homestead Book Co.

6101 22nd Ave. N.W., Seattle, Wa. 98107, 206-782-4532

ISBN: 0-9610798-0-0

Library of Congress Catalog Card Number: 83-070551

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The authors invite comments on The Mushroom Cultivator as well as
personal experiences concerning mushroom cultivation. Address all mail to
Agarikon Press.

Azureus, Skye, and LaDena

TABLE OF CONTENTS

FOREWORD by Dr. Andrew Weil xi

PREFACE xii

I. INTRODUCTION TO MUSHROOM CULTURE 1

An Overview of Techniques for Mushroom Cultivation . 3

Mushrooms and Mushroom Culture 4

The Mushroom Life Cycle 6

II. STERILE TECHNIQUE AND AGAR CULTURE 15

Design and Construction of a Sterile Laboratory 16

Preparation of Agar Media 19

Starting A Culture from Spores 23

Taking a Spore Print 23

Techniques for Spore Germination 24

Characteristics of the Mushroom Mycelium 25

Ramifications of Multispore Culture 25

Sectoring: Strain Selection and Development . 31

Stock Cultures: Methods For Preserving Mushroom Strains 37

III. GRAIN CULTURE 41

The Development of Grain Spawn . . 42

Preparation of Grain Spawn 45

Spawn Formulas 46

Inoculation of Sterilized Grain from Agar Media 48

Inoculation of Sterilized Grain from Grain Masters » • 49

Alternative Spawn Media 54

Liquid Inoculation Techniques 55

Incubation of Spawn 57

IV. THE MUSHROOM GROWING ROOM 61

Structure and Growing Systems 62

Structure 63

Shelves . . . 64

Trays 65

Environmental Control Systems ...... 66

Fresh Air * 66

Fans 68

Air Ducting 70

Filters 70

Exhaust Vents 72

Heating - 73

Cooling

Humidification

Thermostats and Humidistats

Lighting

Environmental Monitoring Equipment

V. COMPOST PREPARATION

Phase I Composting

Basic Raw Materials

Supplements

Formulas

Ammonia

Carbon:Nitrogen Ratio

WaterAir

Pre-Wetting

Building the Pile

Turning

Temperature

Long Composting

Short Composting

Synthetic Compost Procedure

Composting Tools

Characteristics of the Compost at Filling . . . .

Supplementation at Filling

Phase II Composting

Basic Air Requirements

Phase II Room Design

Filling Procedures ,

Depth of Fill

Phase II Procedures: T rays or Shelves

Phase II in Bulk

Bulk Room Design Features

Bulk Room Filling Procedures

Bulk Room Phase II Program

Testing for Ammonia

Aspect of the Finished Compost

Alternative Composts and Composting Procedures

Sugar Cane Bagasse Compost

The Five Day Express Composting Method . .

VI. NON-COMPOSTED SUBSTRATES

Natural Culture

Wood Based Substrates

74
74
74

79
81
82
83

99
100
101
102
104
104

104

105

106
106
106

109

110
114

Straw . 117

VII. SPAWNING AND SPAWN RUNNING IN BULK SUBSTRATES 121

Moisture Content 122

Substrate Temperature 122

Dry Weight of Substrate 122

Duration of Spawn Run 124

Spawning Methods .... ... 124

Environmental Conditions 125

Super Spawning 126

Supplementation at Spawning 126

Supplementation at Casing 126

VIII. THE CASING LAYER 127

Function . ^ ■ 128

Properties ....... 129

Materials 130

Formulas and Preparation ■ 132

Application 133

Casing Colonization 135

Casing Moisture and Mycelial Appearance 137

IX. STRATEGIES FOR MUSHROOM FORMATION (PINHEAD INITIATION) 139

Basic Pinning Strategy 140

Primordia Formation Procedures 141

The Relationship Between Primordia Formation and Yield 146

The Influence of Light on Pinhead Initiation 147

X. ENVIRONMENTAL FACTORS: SUSTAINING THE MUSHROOM CROP 149

Temperature - 130

Flushing Pattern 130

Air Movement 132

Watering 134

Harvesting - 135

Preserving Mushrooms 156

XL GROWING PARAMETERS FOR VARIOUS MUSHROOM SPECIES 159

Agaricus bitorquis 161

Agaricus brunnescens * 164

Coprinus comatus 168

Flammulina velutipes - • 172

Lentinus edodes 176

Lepista nuda • 130

Panaeolus cyanescens ■ ■ 133

Panaeolus subbalteatus 136

Pleurotus ostreatus (Type Variety) .
Pleurotus ostreatus (Florida Variety)
Psilocybe cubensis

Psilocybe cyanescens

Psilocybe mexicana

Psilocybe tampanensis

Stropharia rugoso-annulata . . . .

Volvariella volvacea

XII. CULTIVATION PROBLEMS AND THEIR SOLUTIONS:

A TROUBLE SHOOTING GUIDE 217

Sterile Technique .219

Agar Culture 219

Grain Culture 220

Compost Preparation 223

Phase I 223

Phase II 224

Spawn Running 226

Case Running 227

Mushroom Formation and Development 229

Pinhead Initiation 229

Cropping 231

XIII. THE CONTAMINANTS OF MUSHROOM CULTURE:

IDENTIFICATION AND CONTROL 233

A Key to the Common Contaminants of Mushroom Culture 238

Virus (Die-Back Disease) 244

Actinomyces (Firefang) 246

Bacillus (Wet Spot) 248

Pseudomonas (Bacterial Blotch & Pit) 252

Streptomyces (Firefang) 255

Alternaria (Black Mold) 257

Aspergillus (Green Mold) 259

Botrytis (Brown Mold) 262

Chaetomium (Olive Green Mold) 264

Chrysosporium (Yellow Mold) 266

Cladosporium (Dark Green Mold) 268

Coprinus (Inky Cap) 270

Cryptococcus (Cream Colored Yeast) 273

Dactylium (Cobweb Mold) 275

Doratormyces (Black Whisker Mold) , . 277

Epicoccum (Yellow Mold) 279

Fusarium (Pink Mold)

Geotrichum (Lipstick Mold)

Humicola (Gray Mold)

Monilia (White Flour Mold)

Mucor (Black Pin Mold)

Mycelia Sterilia (White Mold)

Mycogone (Wet Bubble)

Neurospora (Pink Mold

Papulospora (Brown Plaster Mold)

Penicillium (Bluish Green Mold)

Rhizopus (Black Pin Mold)

Scopulariopsis (White Plaster Mold)

Sepedonium (White or Yellow Mold) ....

Torula (Black Yeast)

Trichoderma (Forest Green Mold)

Trichothecium (Pink Mold)

Verticillium (Dry Bubble)

XIV. THE PESTS OF MUSHROOM CULTURE

Mushroom Flies .....

Fly Control Measures

Sciarid Fly

Phorid Fly

Cecid Fly

Mites

Nematodes (Eelworms)

XV. MUSHROOM GENETICS

Reproductive Strategies

Implications for Culture Work

APPENDICES

I. Medicinal Properties of Mushrooms

II. Laminar Flow Systems

III. The Effect of Bacteria and Other Microorganisms on Fruiting

IV. The Use of Mushroom Extracts to Induce Fruiting

V. Data Collection and Environmental Monitoring Records

VI. Analyses of Basic Materials Used in Substrate Preparation . .

VII. Resources For Mushroom Growing Equipment and Supplies

VIII. English to Metric Conversion Tables

GLOSSARY

BIBLIOGRAPHY

281

284

286

288

290

292

294

296

298

300

302

304

306

308

310

313

315

319

320

321
323
325
328
331

333

336

338

343

345

347

253

357

359

369

384

386

389

397

INDEX r . >

PHOTOGRAPHY AND ILLUSTRATION CREDITS
ACKNOWLEDGEMENTS

409

414

415

E ver since French growers pioneered the cultivation of the common Agaricus more than two
hundred years ago, mushroom cultivation in the Western world has been a mysterious art. Pro-
fessional cultivators, fearful of competition, have guarded their techniques as trade secrets, sharing
them only with closest associates, never with amateurs. The difficulty of domesticating mushrooms
adds to the mystery: they are just harder to grow than flowering plants. Some species refuse to grow
at all under artificial conditions; many more refuse to fruit; and even the familiar Agaricus of super-
markets demands a level of care and attention to detail much beyond the scope of ordinary garden-
ing and agriculture.

In the past ten years, interest in mushrooms has literally mushroomed in America. For the first
time in history the English-speaking world is flooded with good field guides to the higher fungi, and
significant numbers of people are learning to collect and eat choice wild species. In the United
States and Canada mushroom conferences and forays attract more and more participants. Culti-
vated forms of species other than the common Agaricus have begun to appear in specialty shops
and even supermarkets.

The reasons for this dramatic change in a traditionally mycophobic part of the world may never
be known. I have been fascinated with mushrooms as symbols of the unconscious mind and think
their growing popularity here is a hopeful sign of progress in the revolution of consciousness that
began in the 1 960s. A more specific reason may be the rediscovery of psychedelic mushrooms—
the Psilocybes and their allies— which have thoroughly invaded American society in recent years.

The possibility of collecting wild psychoactive mushrooms in many parts of North America has
motivated thousands of people to buy field guides and attend mushroom conferences. The possibil-
ity of growing Psilocybe cubensis at home, one of the easier species to cultivate, has made many
people eager to learn the art of mushroom production. As they pursue their hobby, fans of
Psilocybes often find their interest in mushrooms broadening to include other genera that boast
nonpsychoactive but delicious edible species. Other mycophiles, uninterested in altered states of
consciousness, have grown so fond of some edible species as to want better access to them than
foraying in the wild provides. The result has been a demand from a variety of amateurs for the trade
secrets of professional cultivators.

The book you are about to read is a milestone in the new awareness of mushrooms. THE
MUSHROOM CULTIVATOR by Paul Stamets and Jeff Chilton is easily the best source of informa-
tion on growing mushrooms at home. Both authors are experts on the higher fungi, on their techni-
cal aspects as well as the practical methods of working with the most interesting species. Paul
Stamets is a recognized authority on the Psilocybes and their relatives; Jeff Chilton has been a pro-
fessional consultant to large-scale, commercial producers of the common Agaricus and the once-
exotic shiitake of Japan and China. Together they have organized a number of successful mush-
room conferences in the Pacific Northwest and have championed the cause of growing at home.

Unlike experts of the past (and some of the present), they are willing and ready to share their know-
ledge and practical information with all lovers of mushrooms, whether they are amateurs or profes-
sionals, devotees of Psilocybe or of Pleurolus.

THE MUSHROOM CULTVATOR is indeed “A Practical Guide to Growing Mushrooms at
Home,” as its subtitle indicates. It covers every aspect of the subject in a readable style and in suffi-
cient detail to enable both rank amateurs and serious mycologists to succeed at growing the mush-
rooms they like. By including a wealth of excellent illustrations, information on obtaining equipment
and supplies, and step-by-step directions for every procedure, from starting spore cultures to har-
vesting fruiting bodies to dealing with contaminants and pests, the authors demystify the art of
mushroom cultivation and put mastery of it within everyone’s reach. It is a pleasure to introduce this
fine book. If you have been searching for information on this topic, you will find it to be all that you
have been looking for and more.

Andrew Weil, M.D., F.L.S.

T he use of mushrooms as food crosses all cultural boundaries. Highly prized by the
Greeks, mushroom consumption in European nations has deep traditional roots. The
Agari, a pre-Scythian people from Samartia (now Poland and the western Soviet Union), held
mushrooms in high esteem and used them medicinally. The early Greeks held a similar
fascination for fungi and apparently worked them into their religious rituals, even to the extent
that to discuss the use of these sacraments violated strong taboos. For thousands of years, the
Chinese and Japanese have prized a variety of mushroom species for their beneficial proper-,
ties. In the New World, the Aztec and Mazatec Indians of Mexico used mushrooms for both
their healing and divining properties. Clearly, mushrooms have played a significant role in the
course of human cultures worldwide.

Although the Japanese have cultivated the Shiitake mushroom for two thousand years,
the earliest record of European mushroom cultivation was in the 17th century when an
agronomist to Louis XIV, Olivier de Serres, retrieved wild specimens and implanted mush-
room mycelium in prepared substrates. In those times mushroom growing was a small scale
outdoor activity practiced by the rural populace. Materials in which mushrooms grew naturally
were collected and concentrated into prepared beds. These beds were cropped and then used
to start new beds. As demand increased and new methods improved yields, mushroom grow-
ing developed into a large scale commercial business complete with computer controlled in-
door environments and scientifically formulated substrates. Spawn with which to plant
prepared beds, initially gathered in nature, became standardized as sterile culture techniques
were perfected.

It is now known that many of the mushrooms presently under cultivation rank above all
vegetable and legumes (except soybeans) in protein content, and have significant levels of B
and C vitamins and are low in fat. Research has shown that certain cultivated mushrooms
reduce serum cholesterol, inhibit tumors, stimulate interferon production and possess antiviral
properties. It is no surprise, therefore, that as food plants were developed into cultivars, mush-
rooms were among those selected.

Discovering the methods most successful for mushroom cultivation has been a long and
arduous task, evolving from the experience of lifetimes of research. As mushroom growing
expanded from the realm of home cultivators to that of a multimillion dollar industry, it is not
surprising that growers became more secretive about their methods. For prospective home
cultivators, finding appropriate information has become increasingly difficult. As a result, the
number of small growers decreased and home cultivation became a rare enterprise.

The Mushroom Cultivator is written expressly for the home cultivator and is without bias
against any group of interested growers. For the first time, information previously unavailable
to the general public is presented in a clear and easy to understand fashion. The book reflects
not only the work of the authors but also the cumulative knowledge gained through countless

trials by mushrooms growers and researchers. It is the sincere hope of the authors that this
work will re-open the door to the fascinating world of mushroom culture. The Mushroom
Cultivator is dedicated to this goal as we pursue the Art and Science of mushroom cultivation,.

Introduction to Mushroom Culture/ 1

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2/The Mushroom Cultivator

STERILIZATION AND POURING
OF AGAR MEDIUM

PROPAGATION OF PURE CULTURES

STERILIZATION
OF CRAIN MEDIA

germination of spores
AND ISOLATION OF MUSHROOM
MYCELIUM FROM CONTAMINANTS

INOCULATION OF CRAIN

INOCULATION ONTO WOOD DOWELLS

INOCULATION
OF SPAWN

\ LAYING OUT OF SPAWN ON TRAY / Jggp
CASING WITH SOIL-LIKE MIXTURE / ^ ^l|p2s§|

TRAY CULTURE

BAG

CULTURE

Sallkr'

INOCULATION OF BULK SUBSTRATE

IT Li

B\ m

RACK

CULTURE

COLUMN

CULTURE

PLUGGING LOGS

LOG CULTURE

MOUND
(BED) CULTURE

Figure 1 Diagram illustrating overview of general techniques for the cultivation of
mushrooms.

Introduction to Mushroom Culturs/3

AN OVERVIEW OF TECHNIQUES
FOR MUSHROOM CULTIVATION

T echniques for cultivating mushrooms, whatever the species, follow the same basic pattern.

Whereas two species may differ in temperature requirements, pH preferences or the substrate
on which they grow, the steps leading to fruiting are essentially the same. They can be summarized
as follows:

1. Preparation and pouring of agar media into petri dishes.

2. Germination of spores and isolation of pure mushroom mycelium.

3. Expansion of mycelial mass on agar media.

4. Preparation of grain media.

5. Inoculation of grain media with pure mycelium grown on agar media.

6. Incubation of inoculated grain media (spawn).

7. A. Laying out grain spawn onto trays,
or

B. Inoculation of grain spawn into bulk substrates.

8. Casing— covering of substrate with a moist mixture of peat and other materials.

9. Initiation— lowering temperature, increasing humidity to 95%, increasing air circulation,
decreasing carbon dioxide and/or introducing light.

10. Cropping— maintaining temperature, lowering humidity to 85-92%, maintaining air cir-
culation, carbon dioxide and/or light levels.

With many species moderate crops can be produced on cased grain cultures. Or, the cultivator
can go one step further and inoculate compost, straw or wood. In either case, the fruiting of mush-
rooms requires a high humidity environment that can be readily controlled. Without proper mois-
ture, mushrooms don’t grow.

In the subsequent chapters standard methods for germinating spores are discussed, followed by
techniques for growing mycelium on agar, producing grain and/ or bran “spawn” preparing com-
posted and non-composted substrates, spawn running, casing and pinhead formation. With this last
step the methods for fruiting various species diverge and techniques specific to each mushroom are
individually outlined. A trouble-shooting guide helps cultivators identify and solve problems that are
commonly encountered. This is followed by a thorough analysis of the contaminants and pests of
mushroom culture and a chapter explaining the nature of mushroom genetics. In all, the book is a
system of knowledge that integrates the various techniques developed by commercial growers
worldwide and makes the cultivation of mushrooms at home a practical endeavor.

4/The Mushroom Cultivator

Mushrooms inspire awe in those encountering them. They seem different. Neither plant-like
nor animal-like, mushrooms have a texture, appearance and manner of growth all their own. Mush-
rooms represent a small branch in the evolution of the fungal kingdom Eumycota and are common-
ly known as the “fleshy fungi”. In fact, fungi are non-photosynthetic organisms that evolved from
algae. The primary role of fungi in the ecosystem is decomposition, one organism in a succession
of microbes that break down dead organic matter. And although tens of thousands of fungi are
know, mushrooms constitute only a small fraction, amounting to a few thousand species.

Regardless of the species, several steps are universal to the cultivation of all mushrooms. Not
surprisingly, these initial steps directly reflect the life cycle of the mushroom. The role of the culti-
vator is to isolate a particular mushroom species from the highly competitive natural world and im-
plant it in an environment that gives the mushroom plant a distinct advantage over competing
organisms. The three major steps in the growing of mushrooms parallel three phases in their life cy-
cle. They are:

1. Spore collection, spore germination and isolation of mycelium; or tissue cloning.

2. Preparation of inoculum by the expansion of mycelial mass on enriched agar media and
then on grain. Implantation of grain spawn into composted and uncomposted substrates or
the use of grain as a fruiting substrate.

3. Fruitbody (mushroom) initiation and development.

Having a basic understanding of the mushroom life cycle greatly aids the learning of techniques
essential to cultivation.

Mushrooms are the fruit of the mushroom plant, the mycelium. A mycelium is a vast network
of interconnected cells that permeates the ground and lives perenially. This resident mycelium only
produces fruitbodies, what are commonly called mushrooms, under optimum conditions of tem-
perature, humidity and nutrition. For the most part, the parent mycelium has but one recourse for
insuring the survival of the species: to release enormous numbers of spores. This is accomplished
through the generation of mushrooms.

In the life cycle of the mushroom plant, the fruitbody occurs briefly. The mycelial network can
sit dormant for months, sometimes years and may only produce a single flush of mushrooms. Dur-
ing those few weeks of fruiting, the mycelium is in a frenzied state of growth, amassing nutrients and
forming dense ball-like masses called primorida that eventually enlarge into the towering mush-
room structure. The gills first develop from the tissue on the underside of the cap, appearing as
folds, then becoming blunt ridges and eventually extending into flat, vertically aligned plates. These
efficiently arranged symmetrical gills are populated with spore producing cells called basidia.

From a structural point of view, the mushroom is an efficient reproductive body. The cap acts
as a domed shield protecting the underlying gills from the damaging effects of rain, wind and sun.
Covering the gills in many species is a well developed layer of tissue called the partial veil which
extends from the cap margin to the stem. Spores start falling from the gills just before the partial veil
tears. After the partial veil has fallen, spores are projected from the gills in ever increasing numbers.

Introduction to Mushroom Culture/5

6/The Mushroom Cultivator

The cap is supported by a pillar-like stem that elevates the gills above ground where the spores can
be carried off by the slightest wind currents. Clearly, every part of the mushroom fruitbody is de-
signed to give the spores the best opportunity to mature and spread in an external environment that
is often harsh and drastically fluctuating.

As the mushroom matures, spore production slows and eventually stops. At this time mush-
rooms are in their last hours of life. Soon decay from bacteria and other fungi sets in, reducing the
once majestic mushroom into a soggy mass of fetid tissue that melts into the ground from which it
sprung.

Cultivating mushrooms is one of the best ways to observe the entirety of the Mushroom Life
Cycle. The life cycle first starts with a spore which produces a primary mycelium. When the myce-
lium originating from two spores mates, a secondary mycelium is produced. This mycelium con-
tinues to grow vegetatively. When vegetative mycelium has matured, its cells are capable of a
phenomenal rate of reproduction which culminates in the erection of mushroom fruitbody. This
represents the last functional change and it has become, in effect, tertiary mycelium. These types of
mycelia represent the three major phases in the progression of the mushroom life cycle.

Most mushrooms produce spores that are uninucleate and genetically haploid (IN). This
means each spore contains one nucleus and has half the complement of chromosomes for the
species. Thus spores have a “sex” in that each has to mate with mycelia from another spore type to
be fertile for producing offspring. When spores are first released they are fully inflated “moist” cells
that can easily germinate. Soon they dehydrate, collapsing at their centers and in this phase they can
sit dormant through long periods of dry weather or severe drought. When weather conditions pro-

Figure 3 Scanning electron micrograph Figure 4 Scanning electron micrograph
of Russula spores. of Entoloma spores.

Introduction to Mushroom Culture/7

vide a sufficiently moist environment, the spores rehydrate and fully inflate. Only then is germination
possible.

Spores within an individual species are fairly constant in their shape and structure. However,
many mushroom species differ remarkably in their spore types. Some are smooth and lemon
shaped (in the genus Copelandia, for instance); many are ellipsoid (as in the genus Psilocybe);
while others are highly ornamented and irregularly shaped (such as those in Lactarius or Entoloma).
A feature common to the spores of many mushrooms, particularly the psilocybian species, is the
formation of an apical germ pore.

The germ pore, a circular depression at one end of the spore, is the site of germination from
which a haploid strand of mycelium called a hypha emanates. This hypha continues to grow,
branches and becomes a mycelial network. When two sexually complementary hyphal networks
intercept one another and make contact, cell walls separahjjg the two hyphal systems dissolve and
cytoplasmic and genetic materials are exchanged. ErgjifcgJ not, this is “mushroom sex”. Hence-
forth, all resulting mycelium is binucleate and dik^wp^. This means each cell has two nuclei
and a full complement of chromosomes. With few\^^e||jtjjpns, only mated (dikaryotic) mycelia is
fertile and capable of producing fruitbodies. Typically, dikaryotic mycelia is faster running and more

Psilocybe pelliculosa spores.

8/The Mushroom Cultivator

Figure 6 Scanning electron micrograph of a Psilocybe baeocystis spore germinating.

vigorous than unmated, monokaryotic mycelia. Once a mycelium has entered into the dikaryo-
phase, fruiting can occur shortly thereafter. In Psilocybe cubensis, the time between spore germina-
tion and fruitbody initials can be as brief as two weeks; in some Panaeolus species only a week
transpires before mushrooms appear. Most mushroom species, however, take several weeks or
months before mushrooms can be generated from the time of spore germination.

Cultivators interested in developing new strains by crossing single spore isolates take advantage
of the occurrence of clamp connections to tell whether or not mating has taken place. Clamp
connections are microscopic bridges that protrude from one adjoining cell to another and are only
found in dikaryotic mycelia. Clamps can be readily seen with a light microscope at 100-400X
magnification. Not all species form clamp connections. ( Agaricus brunnescens does not; most all
Psilocybe and Panaeolus species do). In contrast, mycelia resulting from haploid spores lack
clamps. This feature is an invaluable tool for the researcher developing new strains. (For more infor-
mation on breeding strategies, see Chapter XV.)

Two dikaryotic mycelial networks can also grow together, exchange genetic material and form
a new strain. Such an encounter, where two hyphal systems fuse, is known as anastomosis. When
two incompatible colonies of mycelia meet, a zone of inhibited growth frequently forms. On agar
media, this zone of incompatibility is visible to the unaided eye.

Introduction to Mushroom Culture/ 9

I pi

f ■.< • '

Figure 7 Scanning electron micrograph of hyphae emanating from a bed of germinat-

ing Psilocybe cubensis spores.

When a mycelium produces mushrooms, several radical changes in its metabolism occurs. Up
to this point, the mycelium has been growing vegetatively. In the vegetative state, hyphal cells are
amassing nutrients. Curiously, there is a gradual increase in the number of nuclei per cell, some-
times to as many as ten just prior to the formation of mushrooms. Immediately before fruitbodies
form, new cell walls divide the nuclei, reducing their number per cell to an average of two. The high
number of nuclei per cell in pre-generative mycelia seems to be a prerequisite for fruiting in many
mushroom species.

As the gills mature, basidia cells emerge in ever increasing numbers, first appearing as small
bubble-like cells and resembling cobblestones on a street. The basidia are the focal point in the re-
productive phase of the mushroom life cycle. The basidia, however, do not mature all at once. In
the genus Panaeolus for instance, the basidia cells mature regionally, giving the gill surface a
spotted look. The cells giving rise to the basidia are typically binucleate, each nucleus is haploid
(1 N) and the cell is said to be dikaryotic. The composition of the young basidia cells are similar. At a
specific point in time, the two nuclei in the basidium migrate towards one another and merge into a
single diploid (2N) nucleus. This event is known as karyogamy. Soon thereafter, the diploid nu-
cleus undergoes meiosis and typically produces four haploid daughter cells.

10/The Mushroom Cultivator

■ppmpd

1*^

SHI

Sag

§6

•i*

—

Figure 8, 9, & 10 Scanning electron micrographs of the mycelial network of Psilocybe
cubensis. Note hyphal crossings and clamp connections.

Introduction to Mushroom Culture/ 11

On the surface of the basidia, arm-like projections called sterigmatae arise through which
these nuclei then migrate. In most species four spores form at the tips of these projections. The
spores continue to develop until they are forcefully liberated from the basidia and propelled into free
space. The mechanism for spore release has not yet been proven. But, the model most widely ac-
cepted within the mycological community is one where a “gas bubble” forms at the junction of the
spore and the sterigmata. This gas bubble inflates, violently explodes and jettisons the spore into the
cavity between the gills where it is taken away by air currents. Most commonly, sets of opposing
spores are released in this manner. With spore release, the life cycle is completed.

Not all mushroom species have basidia that produce four haploid spores. Agaricus
brunnescens (= Agaricus bisporus), the common button mushroom, has basidia with two diploid
(2N) spores. This means each spore can evolve into a mycelium that is fully capable of producing
mushrooms. Agaricus brunnescens is one example of a diploid bipolar species. Some Copelandian
Panaeoli (the strongly bluing species in the genus Panaeolus) are two spored and have mating
properties similar to Agaricus brunnescens. Other mushrooom species have exclusively three
spored basidia; some have five spored basidia; and a few, like the common Chantarelle, have as
many as eight spores per basidium!

An awareness of the life cycle will greatly aid beginning cultivators in their initial attempts to
cultivate mushrooms. Once a basic understanding of mushroom culture and the life processes of
these organisms is achieved, cultivators can progress to more advanced subjects like genetics, strain
selection and breeding. This wholistic approach increases the depth of one’s understanding and
facilitates development of innovative approaches to mushroom cultivation.

Figure 11, 12 & 13 Scanning electron micrographs showing the development ot the
basidium and spores in Ramaria longispora, a coral fungus.

12/The Mushroom Cultivator

Figure 15a, 15b Scanning electron micrographs showing basidium of Psilocybe
pelliculosa. Note spore/sterigmata junction.

Figure 17 Scanning electron micrograph of the gill surface of Cantharellus cibarius.
Note six and eight spored basidia.

14/The Mushroom Cultivator

Sterile Technique and Agar Culture/ 15

16/The Mushroom Cultivator

T he air we breathe is a living sea of microscopic organisms that ebbs and flows with the slightest
wind currents. Fungi, bacteria, viruses and plants use the atmosphere to carry their offspring to
new environments. These microscopic particles can make sterile technique difficult unless proper
precautions are taken. If one can eliminate or reduce the movement of these organisms in the air,
however, success in sterile technique is assured.

There are five primary sources of contamination in mushroom culture work:

1 . The immediate external environment

2. The culture medium

3. The culturing equipment

4. The cultivator and his or her clothes

5. The mushroom spores or the mycelium

Mushrooms— and all living organisms— are in constant competition for available nutrients. In
creating a sterile environment, the cultivator seeks to give advantage to the mushroom over the
myriad legions of other competitors. Before culture work can begin, the first step is the construction
of an inoculation chamber or sterile laboratory.

The majority of cultivators fail because they do not take the time to construct a laboratory for
sterile work. An afternoon’s work is usually all that is required to convert a walk-in closet, a pantry or
a small storage room into a workable inoculation chamber.

Begin by removing all rugs, curtains and other cloth-like material that can harbor dust and
spores. Thoroughly clean the floors, walls and ceiling with a mild disinfectant. Painting the room
with a high gloss white enamel will make future cleaning easier. Cover windows or any other
sources of potential air leaks with plastic sheeting. On either side of the room’s entrance, using plas-
tic sheeting or other materials, construct an antechamber which serves as an airlock. This acts as a
protective buffer between the laboratory and the outside environment. The chamber should be de-
signed so that the sterile room door is closed while the anteroom is entered. Equip the lab with these
items:

1 . a chair and a sturdy table with a smooth surface

2. a propane torch, an alcohol lamp, a bunsen burner or a butane lighter.

3. a clearly marked spray bottle containing a 10% bleach solution.

4. sterile petri dishes and test tube “slants”.

5. stick-on labels, notebook, ballpoint pen and a permanent marking pen.

6. an agar knife and inoculating loop.

All these items should remain in the laboratory, if any equipment is removed, make sure it is
absolutely clean before being returned to the room.

Sterile Technique and Agar Culture/ 17

A semisterile environment can be established in the laboratory through simple maintenance
depending on the frequency of use. The amount of cleaning necessary will be a function of the
spore load in the external environment. In winter the number of free spores drastically decreases
while in the spring and summer months one sees a remarkable increase. Consequently, more
cleaning is necessary during these peak contamination periods. More importantly, all contaminated
jars and petri dishes should be disposed of in a fashion that poses no risk to the sterile lab.

Once the sterile work room has been constructed, follow a strict and unwavering regimen of
hygiene. The room should be cleaned with a disinfectant, the floors mopped and lastly the room’s
air washed with a fine mist of 1 0% bleach solution. After spraying, the laboratory should not be re-
entered for a minimum of 15 minutes until the suspended particles have settled. A regimen of
cleaning MUST precede every set of inoculations. As a rule, contamination is easier to prevent than
to eliminate after it occurs.

Before going further, a few words of caution are required. Sterile work demands concentration,
attention to detail and a steady hand. Work for reasonable periods of time and not to the point of ex-
haustion. Never leave a lit alcohol lamp or butane torch unattended and be conscious of the fact that
in an airtight space oxygen can soon be depleted.

Some cultivators wage war on contamination to an unhealthy and unnecessary extreme. They
tend to “overkill” their laboratory with toxic fungicides and bacteriocides, exposing themselves to
dangerously mutagenic chemical agents. In one incident a worker entered a room that had just
been heavily sprayed with a phenol based germicide. Because of congestion he could not sense the
danger and minutes later experienced extreme shortness of breath, numbness of the extremities and
convulsions. These symptoms persisted for hours and he did not recover for several days. In yet an-
other instance, a person mounted a short wave ultraviolet light in a glove box and conducted trans-
fers over a period of months with no protection and unaware of the danger. This type of light can
cause skin cancer after prolonged exposure. Other alternatives, posing little or no health hazard,
can just as effectively eliminate contaminants, sometimes more so.

If despite one’s best efforts a high contamination rate persists, several additional measures can
be implemented. The first is inexpensive and simple, utilizing a colloidal suspension of light oil into
the laboratory’s atmosphere; the second involves the construction of a still air gtjjjamber called a
glove box; and the third is moderately expensive, employing high efficierjh^aifflpn filters.

1 . By asperating sterile oil, a cloud of highly viscous droplets is create^A^pe droplets des-
cend they trap airborne contaminant particles. This technique uses triethylene glycol that is
vaporized through a heated wick. Finer and more volatile than mineral oil, triethylene glycol
leaves little or no noticable film layer. However a daily schedule of hygiene maintenance is

still recommended. (A German Firm sells a product called an “aero-disinfector” that utilizes
the low boiling point of tri-ethylene glycol. For information write: Chemische Fabrik Bruno
Vogelmann & Co., Postfach 440, 718 Crailsheim, West Germany. The unit sells for less

than $50.00).

2 A glovebox is an airtight chamber that provides a semisterile still air environment in which
to conduct transfers. Typically, it is constructed of wood, with a sneeze window for viewing

18/The Mushroom Cultivator

and is sometimes equipped with rubber gloves into which the cultivator inserts his hands.
Often, in place of gloves, the front face is covered with a removable cotton cloth that is peri-
odically sterilized. The main advantage of a glove box is that it provides an inexpensive, eas-
ily cleaned area where culture work can take place with little or no air movement.

3. Modern laboratories solve the problem of airborne contamination by installing High Effi-
ciency Particulate Air (HEPA) filters. These filters screeen out all particulates exceeding
0. 1 -0.3 microns in diameter, smaller than the spores of all fungi and practically all bacteria.
HEPA filters are built into what is commonly known as a laminar flow hood. Some sterile
laboratories have an entire wall or ceiling constructed of HEPA filters through which pres-
surized air is forced from the outside. In effect, a positive pressure, sterile environment is
created. Specific data regarding the building and design of laminar flow systems is dis-
cussed in greater detail in Appendix IV.

Some cultivators have few problems with contaminants while working in what seems like the
most primitive conditions. Others encounter pronounced contamination levels and have to invest in
high technology controls. Each circumstance dictates an appropriate counter-measure. Whether
one is a home cultivator or a spawn maker in a commercial laboratory, the problems encountered
are similar, differing not in kind, but in degree.

Sterile Technique and Agar Culture/ 19

Once the sterile laboratory is completed, the next step is the preparation of nutrified agar me-
dia. Derived from seaweed, agar is a solidifying agent similar to but more effective than gelatin.
There are many recipes for producing enriched agar media suitable for mushroom culture. The
standard formulas have been Potatoe Dextrose Agar (PDA) and Malt Extract Agar (MEA) to which
yeast is often added as a nutritional supplement. Many of the mycological journals list agar media
containing peptone or neopeptone, two easily accessed sources of protein for mushroom myce-
lium. Another type of agar media that the authors recommend is a broth made from boiling wheat
or rye kernels which is then supplemented with malt sugar.

If a high rate of contamination from bacteria is experienced, the addition of antibiotics to the
culture media will prevent their growth. Most antibiotics, like streptomycin, are not autoclavable and
must be added to the agar media after sterilization while it is still molten. One antibiotic, gentamycin
sulfate, survives autoclaving and is effective against a broad range of bacteria. Antibiotics should be
used sparingly and only as a temporary control until the sources of bacteria can be eliminated. The
mycelia of some mushroom species are adversely affected by antibiotics.

Dozens of enriched agar media have been used successfully in the cultivation of fungi and
every cultivator develops distinct preferences based on experience. Regardless of the type of agar
medium employed, a major consideration is its pH, a logarithmic scale denoting the level of acidity
or alkalinity in a range from 0 (highly acidic) to 14 (highly basic) with 7 being neutral. Species of
Psilocybe thrive in media balanced between 6. 0-7.0 whereas Agaricus brunnescens and allies
grow better in near neutral media. Most mycelia are fairly tolerant and grow well in the 5. 5-7. 5 pH

20/The Mushroom Cultivator

range One needs to be concerned with exact pH levels only if spores fail to germinate or if mycelial
growth is unusually slow.

What follows are several formulas for the preparation of nutritionally balanced enriched agar
media, any one of which is highly suited for the growth of Agaricus, Pleurotus, Lentinus,
Stropharia, Lepista, Flammulina, Volvariella, Panaeolus and Psilocybe mycelia. Of these the
authors have two preferences: PDY (Potatoe Dextrose Yeast) and MPG (Malt Peptone Grain) agar
media. The addition of ground rye grain or grain extract to whatever media is chosen clearly pro-
motes the growth of strandy mycelium, the kind that is generally preferred for its fast growth.

Choose one formula, mix the ingredients in dry form, place into a flask and add water until one
liter of medium is made.

PDY (Potato Dextrose Yeast) Agar

the filtered, extracted broth from boiling
300 grams of sliced potatoes in 1 liter of
water for 1 hour
10 grams dextrose sugar
2 grams yeast (optional)

20 grams agar

MPG (Malt Peptone Grain) Agar

20 grams tan malt
5 grams ground rye grain
5 grams peptone or neopeptone
2 grams yeast (optional)

20 grams agar

For controlling bacteria, 0. 1 0 grams of 60-80% pure gentamycin sulfate can be added to each
liter of media prior to sterilization. (See Resources in Appendix.)

Water quality — its pH and mineral content — varies from region to region. If living in an area of
questionable water purity, the use of distilled water is advisable. For all practical purposes, however,
tap water can be used without harm to the mushroom mycelium. A time may come when balancing
pH is important— especially at spore germination or in the culture of exotic species. The pH of
media can be altered by adding a drop at a time of 1 molar concentration of hydrochloric acid
(HCL) or sodium hydroxide (NaOH). The medium is thoroughly mixed and then measured using a
pH meter or pH papers. (One molar HCL has a pH of O; one molar NaOH has a pH of 12; and
distilled water has a pH of 7).

After thoroughly mixing these ingredients, sterilize the medium in a pressure cooker for 30 min-
utes at 15 psi. (Pressure cookers are a safe and effective means of sterilizing media provided they
are operated according to the manufacturer’s instructions). A small mouthed vessel is recom-
mended for holding the agar media. If not using a flask specifically manufactured for pouring media,
any narrow necked glass bottle will suffice. Be sure to plug its opening with cotton and cover with

MEA (Malt Extract Agar)

20 grams tan malt
2 grams yeast
20 grams agar

(Avoid dark brewer’s malts which have
become carmellized. The malt that
should be used is a light tan brewer’s
malt which is powdery, not sticky in
form).

Sterile Technique and Agar Culture/21

aluminum foil before inserting into the pressure cooker. The media container should be filled only
to % to % of its capacity.

Place the media filled container into the pressure cooker along with an adequate amount of
water for generating steam. (Usually a Vz inch layer of water at the bottom will do). Seal the cooker
according to the manufacturer’s directions. Place the pressure cooker on a burner and heat until
ample steam is being generated. Allow the steam to vent for 4-5 minutes before closing the stop-
cock. Slowly bring the pressure up to 1 5 psi and maintain for Vz hour. Do not let the temperature of
the cooker exceed 250 °F. or else the sugar in the media will caramelize. Media with caramelized
sugar inhibits mycelial growth and promotes genetic mutations.

A sterilized pot holder or newly laundered cloth should be handy in the sterile lab to aid in re-
moving the media flask from the pressure cooker. While the media is being sterilized, immaculately
clean the laboratory.

The time necessary for sterilization varies at different altitudes. At a constant volume, pressure
and temperature directly correspond (a relationship known as Boyle’s Law). When a certain pres-
sure ( = temperature) is recommended, it is based on a sea level standard. Those cultivating at high-
er elevations must cook at higher pressures to achieve the same sterilization effect. Here are two ab-
breviated charts showing the relationships between temperature and pressure and the changes in
the boiling point of water at various elevations. Increase the amount of pressure over the recom-
mended amount based on the difference of the boiling point at sea level and one’s own altitude. For
example, at 5000 feet the difference in the boiling point of water is approximately 10° F. This
means that the pressure must be increased to 20 psi, 5 psi above the recommended 15 psi sea

level standard, to correspond to a “1 0 0

F. increase

” in temperature. (Actually temperature remains

the same; it is pressure that differs).

Relationship of Pressure and

The Relationship of Altitude to

Temperature at Constant Volume

the Boiling Point of Water

Pressure (psi) ° Fahrenheit

Altitude

Boiling Point ( F.)

212

Sea Level

212

220

1,025

210

228

2,063

208

240

3,1 15

206

250

4,169

204

259

5,225

202

267

6,304

200

7,381

197

8,481

196

9,031

195

Note that the effect achieved from sterilizing at 60 minutes at 1 5 psi is the same as that from 30
minutes at 30 psi. Hence a doubling of pressure reduces sterilization time by one half. Most pres-
sure cookers can not be safely operated at this level unless carefully modified according to the

22/The Mushroom Cultivator

Figure 22 & 23 Pouring agar media into sterile
petri dishes. At left, vertical stack technique.

manufacturer’s recommendations. And some extra time must be allowed for adequate penetration
of steam, especially in densely packed, large autoclaves.

Once sterilized, place the cooker in the laboratory or in a semisterile room and allow the pres-
sure to return to 1 psi before opening. One liter of agar media can generously fill thirty 1 00 x 15
mm. petri dishes. Techniques for pouring vary with the cultivator. If only one or two sleeves of petri
dishes are being prepared, the plates should be laid out side by side on the working surface. If more
than two sleeves are being poured or fable space is limited, pouring the sterile petri dishes in a verti-
cal stack is usually more convenient.

Before pouring, vigorously shake the molten media to evenly distribute its ingredients. Experi-
enced cultivators fill the plates rhythmically and without interruption. Allow the agar media to cool
and solidify before using. Condensation often forms on the inside surface of the upper lid of a petri
dish when the agar media being poured is still at a high temperature. To reduce condensation, one
can wait a period of time before pouring. If the pressure cooker sits for 45 minutes after reaching 1
psi, a liter of liquid media can be poured with little discomfort to unprotected hands.

Two types of cultures can be obtained from a selected mushroom: one from its spores and the
other from living tissue of a mushroom. Either type can produce a viable strain of mycelia. Each
has advantages and disadvantages.

Sterile Technique and Agar Culture/23

A mushroom culture can be started in one of two ways. Most growers start a culture from
spores. The advantage of using spores is that they are viable for weeks to months after the mush-
room has decomposed. The other way of obtaining a culture is to cut a piece of interior tissue from
a live specimen, in effect a clone. Tissue cultures must be taken within a day or two from the time
the mushroom has been picked, after which a healthy clone becomes increasingly difficult to estab-
lish.

Taking a Spore Print

To collect spores, sever the cap from the stem of a fresh, well cleaned mushroom and place it
gills down on a piece of clean white paper or a clean glass surface such as a microscope slide. If a
specimen is partially dried, add a drop or two of water to the cap surface to aid in the release of
spores. To lessen evaporation and disturbance from air currents, place a cup or glass over the
mushroom cap. After a few hours, the spores will have fallen according to the radiating symmetry of
the gills. If the spore print has been taken on paper, cut it out, fold it in half, seal in an airtight con-
tainer and label the print with the date, species and collection number. When using microscope
slides, the spores can be sandwiched between two pieces of glass and taped along the edges to
prevent the entrv of contaminant spores. A spore print carelessly taken or stored can easily become
contaminated, decreasing the chance of acquiring a pure culture.

Figure 24a Taking a spore print on typing paper.

24/The Mushroom Cultivator

Figure 24b Taking a spore print on a sterile petri
dish and on glass microscope slides.

Figure 25 Sterilizing two scalpels speeds up
agar transfer technique.

Agaricus brunnescens, Psilocybe cubensis and many other mushroom species have a partial
veil— a thin layer of tissue extending from the cap margin to the stem. This veil can be an aid in the
procurement of nearly contaminant-free spores. The veil seals the gill from the outside, creating a
semi-sterile chamber from which spores can be removed with little danger of contamination. By
choosing a healthy, young specimen with the veil intact, and then by carefully removing the veil
tissue under aseptic conditions, a nearly pure spore print is obtained. This is the ideal way to start a
multispore culture.

Techniques for Spore Germination

Once a spore print is obtained, mushroom culture can begin. Sterilize an inoculating loop or
scalpel by holding it over the flame of an alcohol lamp or butane torch for five or ten seconds until it
is red hot. (If a butane torch is used, turn it down to the lowest possible setting to minimize air dis-
turbance). Cool the tip by inserting it into the sterile media in a petri dish and scrape some spores off
the print. Transfer the spores by streaking the tip of the transfer tool across the agar surface. A simi-
lar method calls for scraping the spore print above an opened petri dish and allowing them to free-
fall onto the medium. When starting a new culture from spores, it is best to inoculate at least three
media dishes to improve the chances of getting a successful germination. Mycelium started in this
manner is called a multispore culture.

When first produced, spores are moist, inflated cells with a relatively high rate of germination.
As time passes, they dry, collapse at their centers and can not easily germinate. The probability of
germinating dehydrated spores increases by soaking them in sterilized water. For 30 minutes at 1 5
psi, sterilize an eye dropper or similar device (syringe or pipette) and a water filled test tube or

Sterile Technique and Agar Culture/25

25-250 ml. Erlenmeyer flask stopped with cotton and covered with aluminum foil. Carefully touch
some spores onto a scalpel and insert into sterile water. Tightly seal and let stand for 6-12 hours.
After this period draw up several milliliters of this spore solution with the eye dropper, syringe or
pipette and inoculate several plates with one or two drops. Keep in mind that if the original spore
print was taken under unsanitary conditions, this technique just as likely favors contaminant spores
as the spores of mushrooms.

Characteristics of the Mushroom Mycelium

With either method of inoculation, spore germination and any initial stages of contamination
should be evident in three to seven days. Germinating spores are thread-like strands of cells emanat-
ing from a central point of origin. These mycelial strands appear grayish and diffuse at first and soon
become whitish as more hyphae divide, grow and spread through the medium.

The mycelia of most species, particularly Agaricus, Coprinus, Lentinus, Panaeolus and
Psilocybe are grayish to whitish in color. Other mushroom species have variously pigmented my-
celia. Lepista nuda can have a remarkable purplish blue mycelium; Psilocybe tampanensis is often
multi-colored with brownish hues. Keep in mind, however, that color varies with the strain and the
media upon which the mycelium is grown. Another aspect of the mycelial appearance is its type of
growth, whether it is aerial or appressed, cottony or rhizomorphic. Aerial mycelium can be species
related or often it is a function of high humidity. Appressed mycelium can also be a species specific
character or it can be the result of dry conditions. The subject of mycelial types is discussed in
greater detail under the sub-chapter Sectoring. (See Color Photos 1-4).

Once the mushroom mycelium has been identified, sites of germinating spores should be
transferred to new media dishes. In this way the cultivator is selectively isolating mushroom mycelia
and will soon establish a pure culture free of contamination. If contamination appears at the same
time, cut out segments of the emerging mushroom mycelia away from the contaminant colonies.
Since many of the common contaminants are sporulating molds, be careful not to jolt the culture or
to do anything that might spread their spores. And be sure the scalpel is cool before cutting into the
agar media. A hot scalpel causes an explosive burst of vapor which in the microcosm of the petri
dish easily liberates spores of neighboring molds.

Ramifications of Multispore Culture

Muitispore culture is the least difficult method of obtaining a viable if not absolutely pure strain.
In the germination of such a multitude of spores, one in fact creates many strains, some incompati-
ble with others and each potentially different in the manner and degree to which they fruit under arti-
ficial conditions. This mixture of strains can have a limiting effect on total yields, with the less pro-
ductive strains inhibiting the activity of more productive ones. In general, strains created from spores
have a high probability of resembling their parents. If those parents have been domesticated and
fruit well under laboratory conditions, their progeny can be expected to behave similarly. In contrast,

26/The Mushroom Cultivator

Figure 26 Stropharia rugoso-annulata spores germinating.

Figure 27 Psilocybe cubensis mycelium growing from agar wedge, transferred from a
multispore germination. Note two types of mycelial growth.

Sterile Technique and Agar Culture/27

cultures from wild specimens may fruit very poorly in an artificial environment. Just as with wild
plants, strains of wild mushrooms must be selectively developed.

Of the many newly created strains intrinsic to multispore germination, some may be only capa-
ble of vegetative growth. Such mycelia can assimilate nutrients but can not form a mushroom fruit-
body (the product of generative growth). A network of cells coming from a single spore is called a
monokaryon. As a rule, monokaryons are not capable of producing fertile spore-bearing mush-
rooms. When two compatible monokaryons encounter one another and mate, cytoplasmic and ge-
netic material is exchanged. The resultant mycelium is a dikaryon that can produce fertile off-
spring in the form of mushrooms. Branching or networking between different dikaryotic strains is
known as anastomosis. This process of recombination can occur at any stage of the cultivation
process: on agar; on grain; or on bulk substrates. The crossing of different mushroom strains is ana-
logous to the creation of hybrids in horticulture.

Another method for starting cultures is the creation of single spore isolates and is accomplished
by diluting spores in a volume of sterile water. This spore solution is further diluted into larger vol-
umes of sterile water which is in turn used to inoculate media dishes. In this way, cultivators can ob-

Figure 28 Four strains of Psilocybe cubensis mycelium: (clockwise, upper right) Matias
Romero; Misantla; Amazonian; and Palenque.

28/The Mushroom Cultivator

serve individual monokaryons and in a controlled manner institute a mating schedule for the devel-
opment of high yielding strains. For cultivators interested solely in obtaining a viable culture, this
technique is unnecessary and multispore germinations generally suffice. But for those interested in
crossing monokaryotic strains and studying mating characteristics, this method is of great value.
Keep in mind that for every one hundred spores, only an average of one to five germinate. For a
more detailed explanation of strains and strain genetics, see Chapter XV.

The greatest danger of doing concentrated multispore germinations is the increased possibility
of contamination, especially from bacteria. Some bacteria parasitize the cell walls of the mycelium,
while others stimulate spore germination only to be carried upon and to slowly digest the resulting
mycelia. Flence, some strains are inherently unhealthy and tend to be associated with a high percen-
tage of contamination. These infected spores, increase the likelihood of disease spreading to neigh-
boring spores when germination is attempted in such high numberSv

Many fungi, however, have developed a unique symbiotic rsl<mMifj|p with other microorgan-
isms. Some bacteria and yeasts actually stimulate spore germinatli^m^iushrooms that otherwise
are difficult to grow in sterile culture. The spores of Cantharellus cibarius, the common and highly
prized Chantrelle, do not germinate under artificial conditions, resisting the efforts of world’s most
experienced mycologists. Recently, Nils Fries (1979), a Swedish mycologist, discovered that when
activated charcoal and a red yeast, Rhodotorula glutinis (Fres.) Flarrison, were added to the media,
spore germination soon followed. (Activated charcoal is recommended for any mushroom whose
spores do not easily germinate.)

igure 29 Psilocybe cubensis spores infected with rod shaped bacteria.

Sterile Technique and Agar Culture/29

Many growers have reported that certain cultures flourish when a bacterium accidentally con-
taminates or is purposely introduced into a culture. Pseudomonas putida, Bacillus megaterium.
Azotobacter vinelandii and others have all been shown to have stimulatory effects on various mush-
room species— either in the germination of spores, the growth of mycelia or the formation of fruit-
bodies (Curto and Favelli, 1972; Hayes ef al., 1969; Eger, 1972; Urayama, 1 961). Techniques
utilizing these bacteria are discussed in Appendix III. However, most of the contaminants one
encounters in mushroom cultivation, whether they are airborne or intrinsic to the culture, are not
helpful. Bacteria can be the most pernicious of all competitors. A diligent regimen of hygiene, the
use of high efficiency particulate air (HEPA) filters and good laboratory technique all but eliminate
these costly contaminants.

STARTING A CULTURE FROM LIVE TISSUE

Tissue culture is an assured method of preserving the exact genetic character of a living mush-
room. In tissue culture a living specimen is cloned whereas in multispore culture new strains are
created. Tissue cultures must be taken from mushrooms within twenty-four to forty-eight hours of
being picked. If the specimens are several days old, too dry or too mature, a pure culture will be dif-
ficult to isolate. Spores, on the other hand, can be saved over long periods of time.

Since the entire mushroom is composed of compressed mycelia, a viable culture can be ob-
tained from any part of the mushroom fruitbody. The cap, the upper region of the stem and/or the
area where the gill plate joins the underside of the cap are the best locations for excising clean tissue.
Some mushrooms have a thick cuticle overlaying the cap. This skin can be peeled back and a tissue
culture can be taken from the flesh underlying it. Wipe the surface of the mushroom with a cotton
swab soaked in alcohol and remove any dirt or damaged external tissue. Break the mushroom cap
or stem, exposing the interior hyphae. Immediately flame a scalpel until red-hot and cool in a media
filled petri dish. Now cut into the flesh removing a small fragment of tissue. Transfer the tissue frag-
ment to the center of the nutrient filled petri dish as quickly as possible, exposing the tissue and agar
to the open air for a minimal time. Repeat this technique into at least three, preferably five more
dishes. Label each dish with the species, date, type of culture (tissue) and kind of agar medium. If
successful, mycelial growth will be evident in three to seven days.

An overall contamination rate of a 1 0% is one most cultivators can tolerate. In primary cultures
however, especially those isolated from wild specimens, it is not unusual to have a 25% contamina-
tion rate. Diverse and colorful contaminants often appear near to the point of transfer. Their num-
bers depend on the cleanliness of the tissue or spores transferred and the hygienic state of the labor-
atory where the transfers were conducted. In tissue culture, the most commonly encountered con-
taminants are bacteria.

Contamination is a fact of life for every cultivator. Contaminants become a problem when their
populations spiral above tolerable levels, an indication of impending disaster in the laboratory. If a
five, ten or fifteen percent contamination rate is normal for a cultivator and suddenly the contamina-
tion level escalates without an alteration of regimen, then new measures of control should be intro-
duced immediately.

30/The Mushroom Cultivator

Figure 30 Splitting
the mushroom stem
to expose interior
tissue.

r ■

/ A ■

Figure 31 Cutting
into mushroom flesh
with a cooled, flame
sterilized scalpel.

I-

\ ^

Figure 32 Excising
a piece of tissue for
transfer into a petri
dish.

Sterile Technique and Agar Culture/31

Once the tissue shows signs of growth, it should be transferred to yet another media dish. If no
signs of contamination are evident, early transfer is not critical. If sporulating colonies of mold devel-
op adjacent to the growing mycelium, the culture should be promptly isolated. Continue transfer-
ring the mycelium away from the contaminants until a pure strain is established. Obviously, isolating
mycelia from a partially contaminated culture is more difficult than transferring from a pure one. The
mere attempt of isolating mycelia away from a nearby contaminant is fraught with the danger of
spreading its spores. Although undetectable to us, when the rim of a petri dish is lifted external air
rapidly enters and spores become airborne. Therefore, the sooner the cultivator is no longer depen-
dent upon a partially contaminated culture dish, the easier it will be to maintain pure cultures. Keep
in mind that a strain isolated from a contaminated media dish can harbor spores although to the
unaided eye the culture may appear pure. Only when this contaminant laden mycelium is inocu-
lated into sterile grain will these inherent bacteria and molds become evident.

To minimize contamination in the laboratory there are many measures one can undertake. The
physical ones such as the use of HEPA filters, asperated oil and glove boxes have already been dis-
cussed. One’s attitude towards contamination and cleanliness is perhaps more important than the
installation of any piece of equipment. The authors have seen laboratories with high contamination
rates and closets that have had very little. Here are two general guidelines that should help many
first-time cultivators.

1 . Give the first attempt at sterile culture the best effort. Everything should be clean: the lab;
clothes; tools; and especially the cultivator.

2. Once a pure culture has been established, make every attempt to preserve its purity. Save
only the cultures that show no signs of mold and bacteria. Throw away all contaminated
dishes, even though they may only be partially infected.

If failure greets one’s first attempts at mushroom culture, do not despair. Only through practice
and experience will sterile culture techniques become fluent.

Agar culture is but one in a series of steps in the cultivation of mushrooms. By itself, agar media
is impractical for the production of mushrooms. The advantage of its use in mushroom culture is
that mycelial mass can be rapidly multiplied using the smallest fragments of tissue. Since contami-
nants can be readily observed on the flat two dimensional surface of a media filled petri dish, it is
fairly easy to recognize and maintain pure cultures.

As mycelium grows out on a nutrient agar, it can display a remarkable diversity of forms. Some
mycelia are fairly uniform in appearance; others can be polymorphous at first and then suddenly de-
velop into a homogeneous looking mycelia. This is the nature of mushroom mycelia— to constantly
change and evolve.

When a mycelium grows from a single inoculation site and several divergent types appear, it is

32/The Mushroom Cultivator

Figure 33 Bacteria growing from con- Figure 34 Rhizomorphic mycelium,
taminated mushroom mycelium. Note divergent ropey strands.

Figure 35 Intermediate linear type myce- Figure 36 Rhizomorphic mycelia with to-
lium. Note longitudinally radial fine mentose (cottony) sector (of Agaricus
strands (Psilocybe cyanescens mycelium). brunnescens).

Sterile Technique and Agar Culture/33

said to be sectoring. A sector is defined solely in contrast to the surrounding, predominant
mycelia. There are two major classes of mycelial sectors: rhizomorphic (strandy) and tomentose
(cottony). Also, an intermediate type of mycelium occurs which grows linearly (longitudinally radial)
but does not have twisted strands of interwoven hyphae that characterize the rhizomorphic kind.
Rhizomorphic mycelium is more apt to produce primordia. Linear mycelium can also produce
abundant primordia but this usually occurs soon after it forms rhizomorphs. Keep in mind, however,
that characteristics of fruiting mycelium are often species specific and may not conform precisely to
the categories outlined here.

In a dish that is largely covered with a cottony mycelia, a fan of strandy mycelia would be called
a rhizomorphic sector, and vice versa. Sectors are common in mushroom culture and although little
is known as to their cause or function, it is clear that genetics, nutrition and age of the mycelium play
important roles.

According to Stoller (1 962) the growth of fluffy sectors is encouraged by broken and exploded
kernels which increase the availability of starch in the spawn media. Working with Agaricus
brunnescens, Stoller noted that although mycelial growth is faster at high pH levels (7.5) than at
slightly acid pH levels (6.5), sectoring is more frequent. He found that sectors on grain could be re-

Figure 37 Psilocybe cubensis mycelia with cottony and rhizomorphic sectors. Note
that primordia form abundantly on rhizomorphic mycelium but not on the cottony
type.

34/The Mushroom Cultivator

Figure 38 Hyphal aggregates of Agaricus bitorquis forming on malt agar media.

Figure 39 Primordia of Psilocybe cubensis forming on malt agar media.

233

Sterile Technique and Agar Culture/35

duced by avoiding exploded grains (a consequence of excessive water) and buffering the pH to 6.5
using a combination of chalk (precipitated calcium carbonate) and gypsum (calcium sulfate).

Commercial Agaricus cultivators have long noted that the slower growing cottony mycelium is
inferior to the faster growing rhizomorphic mycelium. There is an apparent correlation between cot-
tony mycelia on agar and the later occurrence of “stroma”, a dense mat-like growth of mycelia on
the casing which rarely produces mushrooms. Furthermore, primordia frequently form along gen-
eratively oriented rhizomorphs but rarely on somatically disposed cottony mycelia. It is of interest to
mention that, under a microscope, the hyphae of a rhizomorphic mycelial network are larger and
branch less frequently than those of the cottony network.

Rhizomorphic mycelia run faster, form more primordia and in the final analysis yield more
mushrooms than cottony mycelia. One example of this is illustrated in Fig. 37. A single wedge of
mycelium was transferred to a pefri dish and two distinct mycelial types grew from if. The stringy
sector formed abundant primordia while the cottony sector did not, an event common in agar
culture.

When a mycelium grows old it is said to be senescing. Senescent mycelium, like any aged plant
or animal, is far less vigorous and fertile than its counterpart. In general, a change from rhizo-
morphic to cottony looking mycelium should be a warning that strain degeneration has begun.

If at first a culture is predominantly rhizomorphic, and then it begins to sector, there are several
measures that can be undertaken to promote rhizomorphism and prevent the strain’s degeneration.

1 . Propagate only rhizomorphic sectors and avoid cottony ones.

2. Alter the media regularly using the formulas described herein. Growing a strain on the
same agar formula is not recommended because the nutritional composition of the medium
exerts an selective influence on the ability of the mushroom mycelium to produce digestive
enzymes. By varying the media, the strain's enzyme system remains broadly based and the
mycelium is better suited for survival. Species vary greatly in their preferences. Unless
specific data is available, trial and error is the only recourse.

3. Only grow out the amount of mycelium needed for spawn production and return the strain
to storage when not in use. Do not expect mycelium that has been grown over several years
at optimum temperatures to resemble the primary culture from which it came. After so
many cell divisions and continual transfers, a sub-strain is likely to have been selected out,
one that may distantly resemble the original in both vitality, mycelial appearance and fruit-
ing potential.

4. If efforts to preserve a vital strain fail, re-isolate new substrains from multispore germina-
tions.

5. Another alternative is to continuously experiment with the creation of hybrid strains that are
formed from the mating of dikaryotic mycelia of two genetically distinct parents.
(Experiments with Agaricus brunnescens have shown, however, that most hybrids yield
less than both or one of the contributing strains. A minority of the hybrids resulted in more
productive strains.)

36/The Mushroom Cultivator

Home cultivators can selectively develop mushroom strains by rating mycelia according to sev-
eral characteristics. These characteristics are:

1 . Rhizomorphism — fast growing vegetative mycelium.

2. Purity of the strain— lack of cottony sectors.

3. Cleanliness of the mycelia— lack of associated competitor organisms (bacteria, molds and
mites).

4. Response time to primodia formation conditions.

5. Number of primordia formed.

6. Proportion of primordia formed that grow to maturity.

7. Size, shape and/or color of fruitbodies.

8. Total yield.

9. Disease resistance.

10. C0 2 tolerance/sensitivity.

1 1 . Temperature limits.

12. Ease of harvesting.

Using these characteristics, mushroom breeders can qualitatively judge strains and select ones
over a period of time according to how well they conform to a grower’s preferences.

Figure 40 Mature
stand of Psilocybe
cubensis on malt
agar media.

Sterile Technique and Agar Culture/37

Once a pure strain has been created and isolated, saving it in the form of a “stock culture” is
wise. Stock cultures— or “slants” as they are commonly called— are media filled glass test tubes
which are sterilized and then inoculated with mushroom mycelium. A suitable size for a culture tube
is 20 mm. x 1 00 mm. with a screw cap. Every experienced cultivator maintains a collection of stock
cultures, known as a “species bank”. The species bank is an integral part of the cultivation process.
With it, a cultivator may preserve strains for years.

To prepare slants, first mix any of the agar media formulas discussed earlier in this chapter. Fill
test tubes one third of the way, plug with cotton and cover with aluminum foil or simply screw on the
cap if the tubes are of this type. Sterilize in a pressure cooker for 30 minutes at 15 psi. Allow the
cooker to return to atmospheric pressure and then take it into the sterile room before opening. Re-
move the slants, gently shake them to distribute the liquified media and lay them at a 15-30 degree
angle to cool and solidify.

When ready, inoculate the slants with a fragment of mushroom mycelium. Label each tube
with the date, type of agar, species and strain. Make at least three slants per strain to insure against
loss. Incubate for one week at 75 ° F. (24° C.). Once the mycelia has covered a major portion of
the agar’s surface and appears to be free of contamination, store at 35-40 0 F. (2-4° C.). At these
temperatures, the metabolic activity of most mycelia is lowered to a level where growth and nutrient
absorbtion virtually stops. Ideally one should check the vitality of stored cultures every six months by
removing fragments of mycelium and inoculating more petri dishes. Once the mycelium has
colonized two-thirds of the media dish, select for strandy growth (rhizomorphism) and reinoculate
more slants. Label and store until needed. Often, growing out minicultures is a good way to check a
stored strain’s vitality and fruiting ability.

An excellent method to save cultures is by the buddy system: passing duplicates of each
species or of strains to a cultivator friend. Mushroom strains are more easily lost than one might ex-
pect. Once lost, they may never be recovered.

In most cases, the method described above safely preserves cultures. Avid cultivators, however,
can easily acquire fifty to a hundred strains and having to regularly revitalize them becomes tedious
and time consuming. When a library of cultures has expanded to this point, there are several addi-
tional measures that further extend the life span of stock cultures.

A simple method for preserving cultures over long periods of time calls for the application of a
thin layer of sterile mineral oil over the live mycelium once it has been established in a test tube. The
mineral oil is non-toxic to the mycelium, greatly reduces the mycelium’s metabolism and inhibits
water evaporation from the agar base. The culture is then stored at 37-41 0 F. until needed. In a re-
cent study (Perrin, 1 979), all of the 30 wood inhabiting species stored under mineral oil for 27
years produced a viable culture. To reactivate the strains, slants were first inverted upside down so
the oil would drain off and then incubated at 77 0 F. Within three weeks each slant showed renewed
signs of growth and when subcultured onto agar plates they yielded uncontaminated cultures.

38/The Mushroom Cultivator

Figure 41 Filling test tubes with liquid agar media prior to sterilization in a pressure
cooker.

Sterile Technique and Agar Culture/39

Although a strain may be preserved over the long term using this method, will it be as produc-
tive as when it was first stored? Other studies have concluded that strains saved for more than 5
years under mineral oil showed distinct signs of degeneration while these same strains were just as
productive at 2Vz years as the day they were preserved. Nevertheless, it is not unreasonable to
presume, based on these studies, that cultures can be stored up to two years without serious
impairment to their vitality.

Four other methods of preservation include: the immersion of slants into liquid nitrogen (an ex-
pensive procedure); the inoculation of washed sterilized horse manure/straw compost that is then
kept at 36-38 ° F. (See Chapter V on compost preparation); the inoculation of sawdust/bran media
for wood decomposers (see section in Chapter III on alternative spawn media); or saving spores
aceptically under refrigerated conditions — perhaps the simplest method for home cultivators.

Whatever method is used, remember that the mushroom’s nature is to fruit, sporulate and
evolve. Cultivation techniques should evolve with the mushroom and the cultivator must selectively
isolate and maintain promising strains as they develop. So don’t be too surprised if five years down
the line a stored strain poorly resembles the original in its fruiting potential or form.

Figure 43 Culture slant of healthy mycelium
ready for cool storage.

40/The Mushroom Cultivator

42/The Mushroom Cultivator

|S^4Jushroom spawn is used to inoculate prepared substrates. This inoculum consists of a carrier
L ^ § material fully colonized by mushroom mycelium. The type of carrier varies according to the
mushroom species cultivated, although rye grain is the choice of most spawn makers. The history
of the development of mushroom spawn for Agaricus brunnescens culture illustrates how spawn
production has progressed in the last hundred years.

During the 1 800’s Agaricus growers obtained spawn by gathering concentrations of mycelium
from its natural habitat. To further encourage mycelial growth this “virgin spawn” was supple-
mented with materials similar to those occurring naturally, in this case horse manure. Spent com-
post from prior crops was also used as spawn. This kind of spawn, however, contained many con-
taminants and pests, and yielded few mushrooms. Before serious commercal cultivation could
begin, methods guaranteeing the quality and mass production of the mushroom mycelium had to
be developed.

With the advent of pure culture techniques, propagation of mushroom mycelium by spore
germination or by living tissue completely superseded virgin spawn. Now the grower was assured of
not only a clean inoculum but also a degree of certainty as to the strain itself. Strain selection and de-
velopment was possible for the first time in the history of mushroom culture because high yielding
strains could be preserved on a medium of precise composition. Sterilized, chopped, washed com-
post became the preferred medium for original pure culture spawn and was for years the standard of
the Agaricus industry.

In 1932, Dr. James Sinden patented a new spawn making process using cereal grain as the
mycelial carrier. Since then rye has been the most common grain employed although millet, milo
and wheat have also been used. Sinden’s novel approach set a new standard for spawn making and
forms the basis for most modern spawn production. The distinct advantage of grain spawn is the in-
creased number of inoculation sites. Each individual kernel becomes one such point from which
mycelium can spread. Thus, a liter of rye grain spawn that contains approximately 25,000 kernels
represents a vast improvement over inocula transmitted by coarser materials.

Listed below are cereal grains that can be used to produce spawn. Immediately following this
list is a chart illustrating some of the physical properties important to the spawn maker.

RICE: Utilized by few cultivators. Even when it is balanced to recommended moisture levels,
the kernels tend to clump together owing to the sticky nature of the outer coat.

MILLET: Although having a higher number of inoculation points than rye, it is more difficult to
formulate as spawn. Amycel, a commercial spawn-making company, has successfully devel-
oped a formula and process utilizing millet as their primary spawn medium.

SORGHUM: Has spherical kernels and works relatively well as a spawn medium but it can be
difficult to obtain. Milo, a type of sorghum, has been used for years by the Stoller Spawn
Company.

Grain Culture/43

WHEAT: Works equally well as rye for spawn making and fruitbody production.

WHEAT GRASS and RYE GRASS SEED: Both have many more kernels per gram than
grain. The disadvantage of seed is the tendency to lose its moisture and its inability to sepa-
rate into individual kernels, making it difficult to shake. (Rye grass and wheat grass seed are
widely used to promote sclerotia formation in Psilocybe tampanensis, Psilocybe mexicana
and Psilocybe armandii. Perennial or annual can be used although annual is far cheaper.
See the species parameters for these species in Chapter XI.)

RYE: Its availability, low cost and ability to separate into individual kernels are all features
recommending its use as a spawn and fruiting medium.

THE CEREAL GRAINS AND THEIR PHYSICAL PROPERTIES

(tests run by the authors)

TYPE KERNELS/GRAM GRAMS/100 ML % MOISTURE

COMMERCIAL
FEED RYE

15%

COMMERCIAL
MUSHROOM RYE

13%

ORGANIC
CO-OP RYE

11%

ORGANIC WHEAT

10%

SHORT GRAIN
BROWN RICE

100

26%

LONG GRAIN
BROWN RICE

15%

SORGHUM (MILO)

15%

PERENNIAL WHEAT
GRASS SEED

450

16%

PERENNIAL RYE
GRASS SEED

415

12%

MILLET

166

13%

In a single gram of commercial rye, Secale cereale, there is an estimated cell count of
50,000-1 00,000 bacteria, more than 200,000 actinomyces, 1 2,000 fungi and a large number of
yeasts. To sterilize one gram of grain would require, in effect, the destruction of more than 300,000
contaminants! In a spawn jar containing in excess of a hundred grams of grain, and with the addition
of water, the cell population soars to astronomical figures.

Of all the groups of these organisms, bacteria are the most pernicious. Bacteria can divide
every twenty or so minutes at room temperature. At this rate, a single bacterium multiplies into

44/The Mushroom Cultivator

more than a million cells in less than ten hours. In another ten hours, each one of these bacteria
beget another million cells. If only a small fraction of one percent of these contaminants survive the
sterilization process, they can render grain spawn useless within only a few days.

Most microorganisms are killed in the sterilization process. For liquids, the standard time and
pressure for steam sterilization is 25 minutes at 1 5 psi (250 0 F). For solids such as rye, the sterili-
zation time must be increased to insure that the steam sufficiently penetrates the small air pockets
and structural cavities in the grain. Within these cavities bacteria and other thermo-resistant organ-
isms, partially protected from the effects of steam, have a better chance of enduring a shorter sterili-
zation period than a longer one. Hence, a full hour at 1 5 psi is the minimum time recommended to
sterilize jars of rye grain.

Some shipments of grain contain extraordinarily high levels of bacteria and fungi. Correspond-
ingly the contamination rate on these grains are higher, even after autoclaving and prior to
inoculation. Such grain should be discarded outright and replaced with grain of known quality.

Once the grain has been sterilized, it is presumed all competitors have been neutralized. The
next most probable source of contamination is the air immediately surrounding the jars. As hot jars
cool, they suck in air along with airborne contaminants. If the external spore load is excessively
high, many of these contaminants will be introduced into the grain even before conducting a single
inoculation! In an average room, there are 10,000 particulates exceeding .3 microns (dust, spores,
etc.) per cubic foot while in a “sterile” laboratory there are less than 1 00 per cubic foot. With these
facts in mind, two procedures will lessen the chance of contamination after the spawn jars have been
autoclaved.

1 . If autoclaving grain media outside the laboratory in an unsterile environment (a kitchen, for
instance), be sure to clean the outside of the pressure cooker before bringing it into the
sterile inoculating room.

2. Inoculate the jars as soon as they have cooled to room temperature. Although many
cultivators leave uninoculated jars sitting in pressure cookers overnight, this is not recom-
mended.

The amount of water added to the grain is an important factor contributing to the reproduction
of contaminants. Excessive water in a spawn jar favors the growth of bacteria and other competitors.
In wet grain the mushroom mycelium grows denser and slower. Oversaturated grain kernels
explode during the sterilization process, and with their interiors exposed, the grain is even more
susceptible to contamination. In addition, wet grain permeated with mycelium is difficult to break up
into individual kernels. When such grain comes in contact with a non-sterile medium such as casing
soil or compost, it frequently becomes contaminated. Spawn made with a balanced moisture
content has none of these problems. It easily breaks apart into individual mycelium covered kernels,
insuring a maximum number of inoculation points from which mycelial strands can emerge.

Determining the exact moisture content of grain is not difficult. Once done, the cultivator can
easily calculate a specific moisture content that is optimal for use as spawn. Commercial rye grain,
available through co-ops and feed companies, is 1 1% water by mass, plus or minus 2%. The pre-
cise amount of water locked up in grain can be determined by weighing a sample of 100 grams.

Grain Culture/ 45

Then reweigh the same grain after it has been dried in an oven (250 ° F. for 3 hours) and subtract
this new weight from the original 100 grams. The resultant figure is the percentage of moisture nat-
urally bound within the grain.

The optimum moisture content for grain in the production of spawn is between 49-54%. The
following formulas are based on cereal rye grain, Secale cereale, which usually has a moisture
content of 11%. Some variation should be expected depending on the brand, kernel size, geo-
graphical origin and the way the grain has been stored.

The standard spawn container for the home cultivator is the quart mason jar while the
commercial spawn maker prefers the gallon jar. Wide mouth mason jars have been extensively
used by home cultivators because of several books popularizing fruitbody production in these jars.
Wide mouth jars have been preferred because mushrooms grown in them are easier to harvest than
those in narrow mouth ones. Not only is this method of growing mushrooms outdated, but wide
mouth jars have several disadvantages for spawn production and hence are not recommended.
Narrow mouthed containers have less chance of contaminating from airborne spores because of
their smaller openings and are more suited to use with synthetic filter discs. The purpose of the
spawn container is to temporarily house the incubating mycelium before it is laid out in trays or used
to inoculate bulk substrates. Jars are not well suited as a fruiting container.

Most commercial spawn makers cap their spawn bottles with synthetic filter discs which allow
air penetration and gaseous exchange but not the free passage of contaminating spores. Home
cultivators, on the other hand, have used inverted mason lids which imperfectly seal and allow some

Figure 45 Two jars
of grain media,
before and after
autoclaving using the
above formulas.

46/The Mushroom Cultivator

air exchange. This method works fine under sterile conditions although the degree of filtration is not
guaranteed. The best combination uses filter discs in conjunction with one piece screw top lids
having a %-Vi inch diameter hole drilled into its center and fitting a narrow mouthed autoclavable
container. The authors personally find the regular mouthed Vi gallon mason jar to be ideal. (Note:
These Vi gallon jars are inoculated from quart masters, a technique soon to be discussed). Using
only filter discs on wide mouth jars is not recommended due to the excessive evaporation from the
grain medium.

To produce grain spawn of 48-52% moisture use the formulas outlined below and autoclave in
a pressure cooker for 1 hour at 15-18 psi. Note that considerable variation exists between measur-
ing cups, differing as much as 10% in their volumes. Check the measuring cup with a graduated
cylinder. Once standardized, fashion a “grain scoop” and a “water scoop” from a plastic container
to the proportions specified below.

Spawn Formulas
QUART JARS

1 cup rye grain
2 /3 - 3 A cup water

240 ml. grain
170-200 ml.

l/ 2 GALLON JARS

3 cups rye grain
1 3 A cups water
or

(approximately)

600 ml. rye grain
water 400-460 m [water

Figure 46
Commercial spawn
maker’s autoclave.

Grain Culture/47

Figure 47 Pressure cooker of home culti-
vator.

Figure 48 The rubber tire is a helpful
tool for the spawn laboratory. It is used
to loosen grain spawn.

The above formulas fill a quart or a half gallon jar to nearly 2 A of its capacity after autoclaving. In all
these formulas, chalk (CaC0 3 ) and gypsum (CaS0 4 ) can be added at a rate of 1 -3 parts by weight
per 100 parts of grain (dry weight). The ratio of chalk to gypsum is 1:4. The addition of these
elements to spawn is optional for most species but necessary when growing Agaricus brunnescens.
When these calcium buffers are used, add 10% more water than that listed above.

Once the grain filled jars have been autoclaved, they should be placed in the sterile room and
allowed to cool. Prior to this point, the room and its air should be disinfected, either through the use
of traditional cleaning methods, HEPA filters or both.

Upon removing the warm jars from the pressure cooker or autoclave, shake them to loosen the
grain and to evenly distribute wet and dry kernels. Shaking also prevents the kernels at the bottom
of the jar from clumping.

An excellent tool to help in this procedure is a bald car tire or padded chair. Having been care-
fully cleaned and disinfected, the tire should be mounted in an upright and stable position. The tire
has a perfect surface against which to shake the jars, minimizing discomfort to the hands and reduc-
ing the risk of injury from breakage. The tire will be used at another stage in grain culture, so it
should be cleaned regularly. Paint shakers are employed by commercial spawn makers for this
same purpose but they are inappropriate for the home cultivator. CAUTION: ALWAYS INSPECT
THE JARS FOR CRACKS BEFORE SHAKING.

When the grain jars have returned to room temperature, agar to grain inoculations can com-

48/The Mushroom Cultivator

mence. Once again, good hygiene is of the upmost importance. When transferring mycelium from
agar to grain, another dimension is added in which contaminants can replicate. In agar culture, the
mycelium grows over a flat, two dimensional surface. If contamination is present, it is easily seen. In
grain culture, however, the added dimension of depth comes into play and contaminants become
more elusive, often escaping detection from the most discerning eye. If not noticed, contamination
will be spread when this spawn is used to inoculate more sterilized grain.

Before conducting transfers, take precautions to insure the sterile quality of the inoculation en-
vironment. After cleaning the room, do not jeopardize its cleanliness by wearing soiled clothes. Few
cultivators take into consideration that they are a major source of contamination. In fact, the human
body is in itself a habitat crawling with bacteria, microscopic mites, and resplendent with spores of
plants and fungi.

When satisfied that all these preparatory conditions are in force, the making of spawn can
begin.

inoculation of Sterilized Grain from Agar Media

Select a vigorously growing culture whose mycelium covefs 2§ more than % of the agar s sur-
face. Cultures that have entirely overrun the petri dish shb^AlSjjavoided because contaminants
often enter along the margin of the petri dish. If that outer ^dg^s^own over with mycelium, these
invaders can go undetected. Since this peripheral myceliunM^rtufecome laden with contaminant
spores, any grain inoculated with it would become spoiled.

Flame sterilize a scalpel and cut out a triangular wedge of mycelium covered agar using the
technique described for doing agar-to-agar transfers. With careful, deliberate movements quickly
transfer the wedge to an awaiting jar, exposing the grain for a minimal amount of time. For each
transfer, flame sterilize the scalpel and inoculate wedges of mycelium into as many jars as desired. A
petri dish two thirds covered with mycelium should amply inoculate 6-8 quart jars of grain. (A maxi-
mum of 10-12 jars is possible). The more mycelia transferred, the faster the colonization and the
less chance of contamination. Since these jars become the “master cultures , do everything possi-
ble to guarantee the highest standard of purity.

The authors recommend a ’’double wedge” transfer technique whereby a single triangular
wedge of mycelium is cut in half, both pieces are speared and then inserted into an awaiting jar of
sterilized grain. Jars inoculated with this method grow out far faster than the single wedge transfer
technique.

Loosening the lids prior to inoculation facilitates speedy transfers. As each agar-to-grain transfer
is completed, replace the lid and continue to the next inoculation. Once the set is finished, tightly se-
cure the lids and shake each jar thoroughly to evenly distribute the mycelial wedges. In the course of
shaking, each wedge travels throughout the grain media leaving mycelial fragments adhering to the
grain kernels. If a wedge sticks to the glass, distribution is hampered and spawn running is inhibited.
This problem is usually an indication of agar media that has been too thinly poured or has been
allowed to dehydrate. Once shaken, incubate the spawn jars at the appopriate temperature. (A sec-
ond shaking may be necessary on Day 4 or Day 5). In general, the grain should be fully colonized
with mycelium in seven to ten days.

Grain Culture/ 49

Inoculation of Grain from Grain Masters

Once fully colonized, these grain masters are now used for the further production of grain
spawn in quart or Zz gallon containers. Masters must be transferred within a few days of their full col-
onization; otherwise the myceliated kernels do not break apart easily. A step by step description of
the grain-to-grain transfer technique follows.

1 . Carefully scrutinize each jar for any signs of contamination. Look for such abnormalities as:
heavy growth; regions of sparse, inhibited growth; slimy or wet looking kernels (an indica-
tion of bacteria); exploded kernels with pallid, irregular margins; and any unusual colora-
tions. If in doubt lift the lid and smell the spawn— a sour “rotten apple” or otherwise pun-
gent odor is usually an indication of contamination by bacteria. Jars having this scent should
be discarded. (Sometimes spawn partially contaminated with bacteria can be cased and
fruited). Do NOT use any jar with a suspect appearance for subsequent inoculations.

2. After choosing the best looking spawn masters, break up the grain in each jar by shaking
the jars against a tire or slamming them against the palm of the hand. The grain should
break easily into individual kernels. 'Shake as many masters as needed knowing that each
jar can amply inoculate ten to twelve quart jars or seven to nine half gallon jars.

Once completed, SET THE SPAWN JARS ON A SEPARATE SHELF AND WAIT
TWELVE TO TWENTY-FOUR HOURS BEFORE USING. This waiting period is impor-
tant because some of the spawn may not recover, suffering usually from bacterial contami-
nation. Had these jars been used, the contamination rate would have been multiplied by a
factor of ten

3. Inspect the jars again for signs of contamination. After twelve to twenty-four hours, the my-
celium shows signs of renewed growth.

4. If the masters had been shaken the night before, the inoculations can begin the following
morning or as soon as the receiving jars (G-2) have cooled. Again, wash the lab, be person-
ally clean and wear newly laundered clothes.

Place 10 sterilized grain-filled jars on the work-bench in the sterile room. Loosen each
of the lids so they can be removed with one hand. Gently shake the master jar until the
grain spawn separates into individual kernels. Hold the master in your preferred hand. Re-
move the master’s lid and then with the other hand open the first jar to be inoculated. With a
rolling of the wrist, pour one tenth of the master’s contents into the first jar, replace its lid
and continue to the second, third, fourth jars, until the set is completed. When this first set is
done, firmly secure the lids. Replace the lid on the now empty spawn master jar and put it
aside. Take each newly inoculated jar, and with a combination of rolling and shaking, dis-
tribute the mycelium covered kernels evenly throughout.

5. Incubate at the temperature appropriate for the species being cultivated. In a week the
mycelium should totally permeate the grain. Designated G-2, these jars can be used for fur-
ther inoculations, as spawn for the inoculation of bulk substrates, or as a fruiting medium.

Some species are less aggressive than others. Agaricus brunnescens, for instance, can take up

50/The Mushroom Cultivator

Figure 49 Flaming
the scalpel.

Figure 50 Cutting
two wedges of
mycelium colonized
agar.

Figure 51

Inoculating sterilized

Grain Culture/51

to two and a half weeks to colonize grain while Psilocybe cubensis grows through in a week to ten
days. Here again, the use of the tire as a striking surface can be an aid to shaking. For slower grow-
ing species, a common shaking schedule is on the 5th and 9th days after inoculation. The cultures
should be incubated in a semi-sterile environment at the temperature most appropriate for the
species being cultivated. (See Chapter XI).

After transferring mycelium from agar to grain, further transfers can be conducted from these
grain cultures to even more grain filled jars. A schedule of successive transfers from the first inocu-
lated grain jar, designated G-1 , through two more “generations” of transfers (G-2, G-3 respective-
ly) will result in an exponential expansion of mycelial mass. If for instance, 1 0 jars were inoculated
from an agar grown culture (G-1), they could further inoculate 100 jars (G-2) which in turn could
go into 1 000 jars (G-3). As one can see, it is of critical importance that the first set of spawn masters
be absolutely pure for it may ultimately inoculate as many as 1 ,000 jars! Inoculations beyond the
third generation of transfers are not recommended. Indeed, if a contamination rate above 1 0% is ex-
perienced at the second generation of transfers, then consider G-2 a terminal stage. These cultures
can inoculate bulk substrates or be laid out in trays, cased and fruited.

Grain-to-grain transfers are one of the most efficient methods of spawn making. This method is
preferred by most commercial spawn laboratories specializing in Agaricus culture. They in turn sell
grain spawn that is a second or third transfer to Agaricus farmers who use this to impregnate their
compost. For the creation of large quantities of spawn, the grain-to-grain technique is far superior to
agar-to-grain for both its ease and speed. However, every cultivator must ultimately return to agar
culture in order to maintain the purify of the strain.

Figure 2a Spawn master
ready for transfer.

Figure 52b Spawn master
after shaking.

Figure 52c Inoculating
sterilized grain from spawn
master.

Figure 53 Spawn jar contaminated with Wet Spot bacteria, giving the grain a greasy
appearance and emitting a sour odor.

Grain Culture/53

Figure 55 Diagramatic expansion of mycelial mass using grain-to-grain transfer tech-
nique. One petri dish can inoculate 10 spawn jars (G-1) which in turn can be used to
inoculate 100 more jars (G-2) and eventually 1000 jars of spawn (G-3) provided the cul-
ture remains pure.

54/The Mushroom Cultivator

Some mushroom species do not grow well on grain and are better suited to alternative spawn
media. Other mushrooms are grown on substrates incompatible with grain spawn. For example,
sawdust and bran are the preferred spawn materials for the cultivations of wood inhabitors such as
Lentinus edodes and Flammulina velutipes. Another spawn media has a perlite bran base. Perlite is
vitreous rock, heated to 1000°F. and exploded like popcorn. The thin flakes of bran are readily
sterilized while the perlite gives the medium its structure. The recipes are:

Sawdust/Bran Spawn

4 parts sawdust (hardwood)

1 part bran (rice or wheat)

Soak the sawdust in water for a least
twenty four hours, allow to drain and then
thoroughly mix in the bran. If the mixture has
the proper moisture content, a firm squeeze
results in a few drops between the fingers. Fill
the material firmly to the neck of the spawn
container (wide mouth). Japanese spawn
makers bore a Zi inch diameter hole down
the center of the media into which they later
insert their inoculum. Sterilize for 60-90 min-
utes at 15 psi. Once cooled, inoculate from
agar media, liquid emulsion, or grain. A fully
grown bottle of sawdust bran spawn can also
be used for further inoculations.

Perlite Spawn

1 20 milliliters water
40 grams perlite
50 grams wheat bran
6 grams gypsum (calcium sulfate)

1 .5 grams calcium carbonate
Screen the perlite to remove the fine
powder and particulates. Fill the container
(small mouth) with the dry ingredients and
mix well. Add the water and continue mixing
until the ingredients are thoroughly mois-
tened. Sterilize for one hour at 1 5 psi. Inocu-
late from agar media or liquid emulsion.

Figure 56 Mycelium running
through sawdust/bran spawn.

Grain Culture/55

A highly effective technique for inoculating grain utilizes the suspension of fragmented mush-
room mycelia in sterile water. This mycelium enriched solution, containing hundreds of minute
cellular chains, is then injected into a jar of sterilized grain. As this water seeps down through the
grain, mycelial fragments are evenly distributed, each one of which becomes a point of inoculation.
For several days little or no sign of growth may be apparent. On the fourth to fifth day after injection,
given optimum incubation temperatures, sites of actively growing mycelium become visible. In a
matter of hours, these zones enlarge and the grain soon becomes engulfed with mycelium. Using
the liquid inoculation technique eliminates the need for repeated shaking, and a single plate of
mycelium can inoculate up to 1 00 jars, more than ten times the number inoculated by the tradition-
al transfer method. There are several ways to suspend mycelium in water, two of which are de-
scribed here.

The first method is quite simple. Using an autoclaved glass syringe, inject 30-50 ml. of steri-
lized water into a healthy culture. Then scrape the surface of the mycelial mat, drawing up as many
fragments of mycelium as possible. As little as 5 ml. of mycelial suspension adequately inoculates a
quart jar of grain.

The second method incoporates a blender with an autoclavable container-stirrer assembly.
(Several companies sell aluminum and stainless steel units specifically manufactured for liquid cul-
ture techniques— refer to the sources listed in the Appendix). Fill with water until two thirds to three
quarters full, cover with aluminum foil (if a tight fitting metal top is not handy), sterilize and allow to
cool to room temperature.

Under aseptic conditions, insert an entire agar culture of vigorously grown mycelium into the
sterilized stirrer by cutting it into four quadrants or into narrow strips. Because many contaminants
appear along the outer periphery of a culture dish, it is recommended that these regions not be
used. Place all four quadrants or mycelial strips into the liquid. Turn on the blender at high speed for
no longer than 5 seconds. (Longer stirring times result not in the fragmentation of cell chains but in
the fracturing of individual cells. Such suspensions are inviable). Draw up 5-1 0 ml. of the mycelium
concentrate and inoculate an awaiting grain jar.

A further improvement on this technique calls for a 10:1 dilution of the concentrated mycelial
solution. Inject 50 ml. of mycelial suspension into four vessels containing 450 ml. of sterilized
water. Narrow mouth quart mason jars are well suited to this technique. Gently shaking each jar will
help evenly distribute the mycelium. Next incoluate the grain jars with 10-15 milliliters of the diluted
solution. This method results in an exponential increase of liquid inoculum with the water acting as a
vehicle for carrying the mycelial fragments deep into the grain filled jar. This is only one technique
using water suspended fragments of mycelium. Undoubtedly, there will be further improvements as
mycophiles experiment and develop their own techniques.

When using metal lids a small 1 -2 mm. hole can be drilled and then covered with tape. When
the sterilized containers are to be inoculated, remove the tape, insert the needle of syringe, inject the
suspension of mycelia and replace the tape. In this way, the aperture through which the inoculation

Figure 57 Drawing up mycelium from culture dish with syringe.

Figure 58 Syringe inoculation of sterilized grain.

Figure 59 Eberbach container manufactured for liquid culture. Note bolt covering
inoculation hole.

F igure 60 Drawing up liquid inoculum.

Grain Culture/57

— : —

takes place is of minimal size and is exposed for a only second or two. The chance of airborne con-
tamination is minimized.

The liquid inoculation technique works well provided the cultures selected are free from foreign
spores; otherwise the entire set of jars inoculated from that dish will be lost. The disadvantage with
this method is that there is no opportunity to avoid suspect zones on the culture dish— the water sus-
pends contaminant spores and mycelia alike. If a culture dish is contaminated in one region, a few
jars may be lost via the traditional inoculation method while with the liquid inoculation technique
whole sets of up to one hundred spawn jars would be made useless.

Although mycelial suspensions created in this manner work for many species, the mycelia of
some mushrooms do not survive the stirring process.

INCUBATION OF SPAWN

With each step in the cultivation process, the mycelial mass and its host substrate increases. In
seven days to two weeks after inoculation, the spawn jars should be fully colonized with mushroom
mycelia. The danger here is that, if contamination goes undetected, that mold or bacterium will like-
wise be produced in large quantities. Hence, as time goes by the importance of clean masters be-
comes paramount. By balancing environmental parameters during incubation, especially
temperature, the mycelium is favored.

Once the jars have been inoculated, store them on shelves in a semisterile room whose tem-
perature can be easily controlled. Light and humidity are not important at this time as a sealed jar
should retain its moisture. Air circulation is important only if the incubating jars overheat. In packing
a room tightly with spawn jars, overheating is a danger. Many thermophilic fungi that are inactive at
room temperature flourish at temperatures too high for mushrooms. Herein lies one of the major
problems with rooms having a high density of incubating spawn jars. If possible, some provisions
should be made to prevent temperature stratification in the incubation environment.

The major factor influencing the rate of mycelial growth is temperature. For every species there
is an optimum temperature at which the rate of mycelial growth is maximized. As a general rule, the
best temperature for vegetative (spawn) growth is several degrees higher than the one most stimula-
tory for fruiting. In Chapter XI, these optimum temperatures and other parameters are listed for
more than a dozen cultivated mushrooms. Yet another factor affecting both growth rate and suscep-
tibility to contamination is moisture content, a subject covered in the previous chapter on grain
culture.

Every day or so inspect the jars and check for the slightest sign of contamination. The most
common are the green molds Penicillium and Aspergillus. If contamination is detected, seal the lid
and remove the infected culture from the laboratory and growing facility. If a jar is suspected to be
contaminated, mark it for future inspection.

Not all discolorations of the grain are de facto contaminants. Mushroom mycelium exudes a
yellowish liquid metabolite that collects as droplets around the myceliated kernels of grain. As the
culture ages and the kernels are digested, more metabolic wastes are secreted. Although this secre-

58/The Mushroom Cultivator

Figure 61 Half gallon jars of spawn incu- Figure 62 Gallon jars incubating in semi-
bating in semisterile environment. sterile environment.

tion is mostly composed of alcohols (ethanol and acetone), in time acids are produced that cause
the lowering of the substrate’s pH. These waste products are favorable to the propagation of bacteria
that thrive in aqueous environments. Small amounts of this fluid do not endanger the culture;
excessive waste fluids (where the culture takes on a yellowish hue) are definitely detrimental. If this
fluid collects in quantities, the mycelium sickens and eventually dies in its own wastes. Such exces-
sive “sweating” is indicative of one or a combination of the following conditions:

1 . Incubation at too high a temperature for the species being cultivated. Note that the tempera-
ture within a spawn jar is several degrees higher than the surrounding air temperature.

2. Over-aging of the cultures; too lengthy an incubation period.

3. Lack of gas exchange, encouraging anaerobic contamination,

Contaminated jars should be sterilized on a weekly basis. Do not dig out moldy cultures unless
they have been autoclaved or if the identity of the contaminant in question is known to be benign.
Several contaminants in mushroom culture are pathogenic to humans, causing a variety of skin dis-
eases and respiratory ailments. (See Chapter XIII on the contmaninants of mushroom culture).
Autoclave contaminated jars for 30 minutes at 15 psi and clean soon after. Many autoclaved jars,
once contaminated, re-contaminate within only a few days if their contents have been not discarded.

Grain Culture/59

Figure 63 Chart showing influence of temperature on the rate of mycelial growth in
Psilocybe cubensis and Psilocybe mexicana. (Adapted from Ames et al., 1958).

If an exceptionally high contamination rate persists, review the possible sources of contamina-
tion, particularly the quality of the master spawn cultures (such as the moisture content of the grain)
and the general hygiene of the immediate environment. Once the cultures have grown through with
mycelium and are of known purity, this spawn can be used to inoculate bulk substrates or can be
layed out in trays, cased and fruited.

60/The Mushroom Cultivator

The Mushroom Crowing Room/61

Figure 64 Small growing room utilizing shelves.

62/The Mushroom Cultivator

M ushroom cultivation was originally an outdoor activity dependent on seasonal conditions.

Substrates were prepared and spawned when outside temperatures and humidity were
favorable. This is still the case with many small scale growers of Volvariella volvacea, Stropharia
rugoso-annulata and Lentinus edodes.

Agaricus cultivators grow solely indoors. Initially, Agaricus growers in France adapted the lime-
stone mines near Paris and in the Loire valley to meet the necessary cultural requirements of that
mushroom. These “caves” were well suited because of their constant temperature and high humid-
ity, essential requirements for mushroom growing. When the first houses designed solely for mush-
rooms were built in the early 1 900’s, temperature and humidity control were the main factors guid
ing their construction.

For the home cultivator, a growing room should be scaled according to the scope of the proj-
ect. The following guidelines supply the information to properly design and equip a growing cham-
ber, basement growing room, outdoor shed or garage.

Figure 65 An insulated plastic greenhouse suitable for mushroom growing.

The Mushroom Growing Room/63

Structure

The basic structure of a mushroom house is made of wood or concrete block with a cement
floor. Because water collects on the floor during the cropping cycle, provisions should be made for
drainage. A wood floor can be covered with a heavy guage plastic. Interior walls, ceilings and ex-
posed wood surfaces should be treated with a marine enamel or epoxy-plastic based paint. A white
color enhances lighting and exposes any contaminating molds.

The most important feature of a growing room is the ability to maintain a constant temperature.
In this respect, insulation is critical. The walls should be insulated with R = 1 1 or R = 19 and the ceil-
ing with R = 30 insulating materials. Fiberglass or styrofoam work well but should be protected
from the high humidities of the growing room to prevent water from saturating them. For this pur-
pose, a 2-4 mil. plastic vapor barrier is placed between the insulation and the interior wall.

An airtight room is an essential feature of the mushroom growing environment, preventing in-
sects and spores from entering as well as giving the cultivator full control over the fresh air supply.
During the construction or modification of the room, all cracks, seams and joints should be carefully
sealed.

Many growers modify existing rooms in their own homes or basements. The main considera-
tion for this approach is to protect the house structure (normally wood) from water damage and to
make the growing chamber airtight. This is accomplished by plastic sheets stapled or taped to the
walls, ceiling and floor, with the seams and adjoining pieces well sealed. If the room is adjacent to an
exterior house wall where a wide temperature fluctuation occurs, condensation may form between
the plastic and the wall. Within these larger structures, a plastic tent or envelope room can be con-

Figure 66 Cultivation of mushrooms in an aquarium.

64/The Mushroom Cultivator

structed. Such a structure can be framed with 2” PVC pipe. The pipe forms a box frame to which
the plastic is attached. This type of growing room should not need insulation because of the air buf-
fer between it and the larger room.

Porches, basements and garages can all be modified in the ways just mentioned. These areas
can also be used with little additional change if the climate of the region is compatible with the
mushroom species being grown. For example, Lentinus edodes, the shiitake mushroom, readily
fruits at 50-60 degrees F. in a garage or basement environment.

The newest innovation in mushroom growing structures is the insulated plastic greenhouse.
The framework is made of galvinized metal pipe bent into a semi-circular shape and mounted at
ground level or on a 3.5 foot side wall. The ends of the walls and the doors are framed with wood.
Heavy plastic (5-6 mil) is stretched over the metal framework to form the inner skin of the room. A
layer of wire mesh is laid over the plastic and functions to hold 3-6 inches of fiberglass insulation in
place. A second plastic sheet covers the insulation and protects it from the weather. The plastic
should be stretched tight and anchored well. These layers are held in place by structural cable span-
ning the top and secured at each side. (See Figure 65). This type of structure, plastic coverings and
plastic fasteners are all available at nursery supply companies. Remember, the design of a mush-
room growing room strives to minimize heat gain and loss.

For people with little or no available space, “mini-culture” in small environmental chambers
may be the most appropriate way to grow mushrooms. Styrofoam ice chests, aquariums and plastic
lined wood or cardboard boxes can all be used successfully. Because of the small volume of sub-
strates contained in one of these chambers, air exchange requirements are minimal. Usually,
enough air is exchanged in opening the chamber for a daily or twice daily misting. Clear, perforated
plastic covering the opening maintains the necessary humidity and the heat can be supplied by the
outer room. Larger chambers can be equipped with heating coils or a light bulb on a rheostat. Both
should be mounted at the base of the chamber. Mini-culture is an excellent and proven way to grow
small quantities of mushrooms for those not having the time or resources to erect larger, more con-
trolled environments.

Shelves

The most common indoor cultivation method is the shelf system. In this system, shelves form a
platform upon which the mushroom growing substrate is placed. The shelf framework consists of
upright posts with cross bars at each level to support the shelf boards. This fixed framework is con-
structed of wood or non-corrosive tubular metal. The shelves should be a preservative-treated soft-
wood. The bottom boards are commonly six inches wide with one inch spaces between them. Side
boards are 6-8 inches high depending on the depth of fill. A standardized design is shown in Figure
67. All shelf boards are placed unattached thereby allowing easy filling, emptying and cleaning.
Agaricus growers fill the shelf house from the bottom up. The shelf boards are stacked at the side of
the room and put into place after each level is completed.

The center pole design (shown in Figure 67) is a simple variation that is less restrictive and
ideally suited for growing in plastic bags. Another alternative is to use metal storage shelves. These

The Mushroom Growing Room/65

Figure 67 Double support and centerpole design shelves. Both shelves are firmly at-
tached to the floor and the ceiling.

units come in a variety of widths and lengths and have the distinct advantage of being impervious to
disease growth. Their use is particularly appropriate for cropping on sterilized substrates in small
containers.

Trays

The development of the tray system in Agaricus culture is largely due to the work of Dr. James
Sinden. In direct contrast to anchored static shelves, trays are individual cropping units that have the
distinct advantage of being mobile. This mobility has made mechanization of commercial cutlivation
possible. Automated tray lines are capable of filling, spawning and casing in less time, with fewer
people and with better quality management.

Whereas in the shelf system all stages of the cultural cycle occur in the same room, the tray sys-
tem utilizes a separate room for Phase II composting. On a commercial tray farm only the Phase II
room is equipped for steaming and high velocity air movement.

A Sinden system tray design is shown in Figure 68. This tray has short legs in the up-position.
During Phase II and spawn running these trays are stacked 15 cm. apart and tightly placed within
the room to fully utilize compost heat. After casing, a wooden spacer is inserted between the trays
for crop management, increasing the space to 25-35 cm. Other tray designs have longer legs in the
down position and higher sideboards to accomodate more compost. These trays are similarly
spaced throughout the cycle. In the growing room, trays can be stacked 3-6 high in evenly spaced
rows. The main considerations for the home cultivator are that the trays can be easily handled and
that they fit the floor space of the room.

The real advantage of the tray system is the ability to fill, spawn and case single units in an unre-

66/The Mushroom Cultivator

Figure 68 Sinden system
tray. The tray can be con-
structed of 1 x 6 or 2 x 6 inch
lumber for bottom and side
boards and 4x4 inch corner
posts. (1 x 8 or 2 x 8 inch side
boards are suggested for
deep fills).

stricted environment outside the actual growing room. The tray system also gives the cultivator
more control over hygiene and improves the efficiency of the operation. Moving trays from room to
room does present contamination possibilities; therefore, the operations room must also be clean
and fly tight for spawning and casing. Because there is no fixed framework in the growing room, it is
easily cleaned and disinfected.

The tray method has many distinct advantages over the mason jar method for home cultivators
preferring to fruit mushrooms on sterilized grain. These advantages are: fewer necessary spawn
containers; fewer aborts due to uncontrolled primordia formation between the glass/ grain interface;
ease of picking and watering; better ratio of surface area to grain depth; and comparatively higher
yields on the first and second flushes. An inexpensive tray is the 3-4 inch deep plant propagation flat
commonly sold for staring seedlings. An example of such a tray is pictured in Fig. 69.

The mushroom growing room is designed to maintain a selected temperature range at high rel-
ative humidities. This is accomplished through adequate insulation and an environmental control
system with provisions for heating, cooling, humidification and air handling.

In the original shelf houses the environment was controlled by a combination of active and
passive means. Fresh air was introduced through adjustable vents running the length of the ceiling
above the center aisle. Heat was supplied by a hot water pipe along the side walls, a foot above
ground level. And humidity was controlled by similarly placed piping carrying live steam. The warm
air rising up the walls in combination with the cool fresh air falling down the center aisle created con-
vection currents for air circulation. Although no longer in general use by Agaricus growers, air
movement based on convection can be similarly designed for small growth chambers where me-
chanical means are inappropriate.

Present day Agaricus farms integrate heating, cooling and humidification equipment into the
air handling system and in this way are able to achieve balanced conditions throughout the growing
room. Figure 73 shows an example of this type of system.

Fresh Air

Filtered fresh air enters the room at the mixing box where it is proportionally regulated with re-

The Mushroom Growing Room/67

Figure 69 Psilocybe cubensis fruiting on
cased grain in a tray.

Figure 70 Psilocybe cubensis fruiting in
pint and a half jars.

Figure 71 Psilocybe cubensis fruiting in
pint jars.

Figure 72 Psilocybe cubensis fruiting in
a plastic lined box.

Figure 73 Standard ventilation system used by Agaricus growers. (After Vedder)

circulated air by a single damper. To prevent leakage during spawn running and pre-pinning, the
damper fits tightly against the fresh air inlet. This allows full recirculation of room air to maintain
even conditions, thereby counteracting temperature and C0 2 stratification. When fresh air is re-
quired, the damper can be adjusted to any setting, including complete closure of the recirculation in-
let. As fresh air is introduced, room air is displaced and evacuated through an exhaust vent or cracks
around the door. Because fresh air is generally at a different temperature than the one required for
the growing room, it must be used judiciously in order to avoid disrupting the growing room envi-
ronment or overworking the heating, cooling and humidication systems. By properly mixing the
fresh outside air and the room air, a balance can be achieved and optimum conditions for mush-
room growth prevail.

Fresh air serves many important functions in mushroom culture, primarily by supplying oxygen
to the growing mushrooms and carrying away C0 2 . Fresh air also facilitates moisture evaporation
from the cropping surface. To determine the exact amount of air needed in a given situation, a
knowledge of the C0 2 requirements for the species being grown is necessary. (See Chapter XI on
growing parameters for various species). The fresh air can also be measured in terms of air changes
per hour, a common way mushroom growers size the fan in the growing room.

Fans

Axial flow and centrifugal fans are the two most commonly used in mushroom houses. Both
fans operate well against high static pressure, which is a measure of the resistance to forced air.
Static pressure is measured in inches of water gauge— the height in inches to which the pressure lifts
a column of water— and is caused by filters, heating and cooling coils or other obstructions to the
free flow of air. Fans are rated in terms of their output, a measurement of cubic feet per minute
(CFM) at varying static pressures (S.P.). When choosing a fan, these two factors must be considered
for proper sizing.

70/The Mushroom Cultivator

substrates. The reason this ratio is so important is that increased amounts of substrate can generate
heat and carbon dioxide beyond the handling capacity of the ventilation system. A large free air
space acts to buffer these changes. Ostensibly, a ventilation system could be matched with a room
having a 3:1 air-to-bed ratio, but it would have to move such a volume of air that evaporation off the
sensitive cropping surface would be uncontrollable and excessive. Growing mushrooms on thin
layers of grain (1 -3 inches), however, produces less CO 2 than growing on 8 inches of compost and
consequently would emit a lower air-to-bed ratio.

Air Ducting

Ducting for the air system is standard inflatable polyethylene tubing, sized to conform to the fan
diameter. If ducting is not available in the correct size, PVC pipe can be substituted. Figures 74,
75 and 76 show different air distribution arrangements and their flow patterns. The ducts run the
length of the room at ceiling level. One is centrally mounted and discharges towards both walls or
directly down the center aisle, whereas the other is wall mounted and is directed across the width of
the room.

The outlet holes in the duct are designed to discharge air at such a velocity that the airstream
reaches the walls and passes down to the floor without directly hitting the top containers. The holes
are spaced so that the boundaries of the adjacent jets meet just before reaching the wall or floor. This
effectively eliminates dead-air pockets. To size and space the outlet holes exactly, two guidelines are
used:

1 . The total surface area of the holes is equal to the cross section of the duct. (The area of a cir-
cle is 2 % times the radius squared, A=n(r) 2 ).

2. The space between the holes is equal to a quarter of the distance between the duct and the
wall or floor.

The discharge of air at velocities sufficient to draw in surrounding room air is called entrain-
ment, a phenomenon that enhances the capacity of the air circulation system. A flow pattern of even
air is then reached that directly benefits the growing mushrooms. The entrainment of air is the goal
of air management in the growing room.

Filters

Fresh air filters are an important part of the ventilation system and contribute to the health of the
crop. Their function is to screen out atmospheric dust particles like smoke, silica, soot and decayed
biological matter. Atmospheric dust also contains spores, bacteria and plant pollen, some of which
are detrimental to mushroom culture. Furthermore, spores and microorganisms originating within
the cropping room can also be spread by air movement. To counteract this danger, some mush-
room farms filter recirculated air as well.

Agaricus growers commonly use high efficiency, extended surface, dry filters. These filters are
of pleated or deep fold design which gives them much more surface area than their frame opening.
They filter out 0.3 micron particles with 90-95% efficiency and 5.0 micron particles with an effi-

Figure 74 Central aisle
outward flow air
circulation pattern.

Figure 75 Central aisle
downflow air circulation
pattern with wall
mounted baseboard
heating.

Figure 76 Wall
mounted duct directing
airflow across the width
or down the sidewall of
the room.

72/The Mushroom Cultivator

MISTING NOZZLE

^ PRE FILTER

.3 MICRON FILTER

Figure 77 Schematic of mixing box and controlled recirculation system in the growing
room.

ciency of 99% at an initial resistance of 0.10 to 0.50 inches of static pressure.

High efficiency particulate air (HEPA) filters are even more efficient than those just described
and are cost effective for the home cultivator. They screen out particulates down to 0. 1 -0.3 microns
with a rated 99.96% to 99.99% efficiency and have a resistance of .75-1 .00 inches of static pres-
sure. HEPA filters are made of a variety of materials, depending on their intended application. Most
HEPA filters operate in environments of up to 80% humidity without disintegration. Special “water-
proof” filters operable in 100% humidity environments can also be purchased at little or no extra
expense. These “waterproof” filters are especially appropriate for use with systems that push recir-
culated air through the filter. This type of system is illustrated in Figure 77. To protect the filters and
prolong their usefulness, a one inch prefilter of open celled foam or fiberglass (of the furnace type) is
installed to remove large particulates.

Exhaust Vents

Exhaust vents are designed to relieve overpressure within the growing room caused by the in-
troduction of fresh air. Without an exit for the air, a back pressure is created that increases resistance
and reduces the CFM of the fan. Small rooms operating with low fresh air requirements can forgo
special exhaust vents and allow the air to escape around the seals of the room entrance, in effect
creating a positive pressure environment. Positive pressure within a room can also be created by

The Mushroom Growing Room/73

undersizing the exhaust vent, which should be no larger than half the size of the fresh air inlet. Free
swinging dampers operating on overpressure are widely employed in the mushroom growing in-
dustry, The outlet should be screened from the inside to prevent the entry of flies.

Heating

bleating systems for cropping rooms can be based on either dry heat or live steam. Dry heat
refers to a heating source that lowers the moisture content of the air as it raises the temperature.
These systems utilize either hot water or steam circulating through a closed system of pipes or
radiator coils. Heating systems can also be simple resistance coils or baseboard electric heaters.
Heat coils are placed in the air circulation system ahead of the fan as shown in Figure 73. Small por-
table space heaters can also be attached to the mixing box or placed on the wall under it. Otherwise,
baseboard heaters can be installed along the length of the side walls and matched with the air circu-
lation design shown in Figure 75.

Heat supplied by live steam has the advantage of keeping the humidity high while raising the
temperature of the room. If regulated correctly, steam can maintain the temperature and relative hu-
midity within the required ranges without drawing upon other sources. Nevertheless, a backup heat
source is advantageous in the event humidity levels become too high. For steam heat to function
properly it should be controlled volumetrically by adjusting a hand valve (rather than simply on and
off). Vaporizers well suited for small growing rooms are available in varying capacities, and can be
fitted with a duct that connects with the air system downstream from the fan and filter.

To avoid high energy consumption and the expense associated with equipment purchase,
operation and maintenance, the growing room should be designed to take full advantage of the heat
generating capabilities of the substrate. This is done by matching the air-to-bed ratio to the type of
substrate. Growing on thin layers of grain can be done with a ratio of 4:1 (or less) whereas compost
demands 5:1 . The influence of the outside climate and its capacity for cooling the growing room
should also be considered. All these factors must be evaluated before a growing environment with
efficient temperature control can be constructed.

Cooling

Commercial farms use cooling coils with cold water or glycol circulating through them. The
coils are placed before the fan as shown in Figure 73 and are supplied by a central chiller or under-
ground tank and well. Other systems use home or industrial refrigeration or air conditioning units
that operate with a compressor and liquid coolant filled coils. These units are positioned to draw in
recirculated as well as fresh air. All these systems share the common trait of drawing warm air over a
colder surface. In doing so, moisture condenses out of the air and in effect dehumidifies the room.

The oldest and most widely practiced method of cooling is through the use of fresh air. Cooling
with fresh air depends upon the weather and the temperature requirements of the species being cul-
tivated. However, its use is the most practical means available to the home cultivator. In climates
with high daily temperatures, fresh air can be shut off or reduced to a minimum during the day and

74/The Mushroom Cultivator

then fully opened at night when temperatures are at their lowest.

Humidification

Most mushroom growers use steam as the principal means of humidification. The steam is in-
jected into the air system duct on the downstream side of the fan and filter. Household vaporizers
are well suited for small growing rooms. They are available in various capacities and can be fitted
with a duct running to the air system. The vaporizer can also be positioned under the mixing box for
steam uptake with the recirculated air. Keep in mind that cold fresh air has much less capacity for
moisture absorbtion and therefore does not mix well with large volumes of steam.

Another method of humidification uses atomizing nozzles to project a fine mist into the air
stream. Large systems have a separate mixing chamber with nozzles mounted to spray the passing
air. In a small room, a single nozzle can be mounted in the center of the duct and aimed to flow with
the air as it exits the fan. (See Figure 77). An appropriately sized nozzle emits 0.5-1 .0 gallons per
hour at 20-30 psi. To prevent the nozzle from plugging up, filters should be incorporated in the
water supply line.

In a third method, air passes through a coarse mesh absorbant material that is saturated with
water. This system is widely used for cooling at nurseries. It is similar in principle to a “swamp
cooler”. In this system (and the water atomizing system), the temperature of the supply water can be
regulated to provide a measure of heating and cooling. Both systems also produce some free water
so provisions must be made for drainage.

Thermostats and Humidistats

In general, thermostats and humidistats are designed to open and close valves in response to
pre-set temperature or humidity limits. The instrument sensors are placed in a moving air stream
representative of room conditions, usually in or near the recirculation inlet. Because these instru-
ments are programmed for either on or off, heat and humidity come in surges. Often this results in
uneven and fluctuating conditions within the room.

The ideal in environmental control is to supply just enough heat and humidity to make up for
losses from the room and to compensate for differences in the fresh air. Modulating thermostats do
this by supplying heat continously in proportion to the deviation from the desired temperature. Posi-
tive control of this sort can also be accomplished by hand valves, alone or in conjunction with
on/ off instruments. Supply line volume is thereby regulated in order to attain an equilibrium. With a
thermostat, this means keeping the supply volume just below the cut-off point.

Lighting

Many cultivated mushrooms require light for pinhead initiation and proper development of the
fruitbody. In fact, such phototropic mushrooms actually twist and turn towards a light source, espe-
cially if it is dim and distant in an otherwise darkened room. Consequently, it is important to equip

The Mushroom Growing Room/75

Figures 78, 79 & 80 Charts
showing the proportions of
spectra in incandescent, fluores-
cent and natural lighting.

»«

S70 S« US

76/The Mushroom Cultivator

Figure 81 An inexpensive hygrometer for
measuring relative humidity.

the growing room with a lighting system that provides even illumination to all areas and levels.

Flourescent light fixtures are the most practical and give the broadest coverage. These fixtures
should be evenly spaced and mounted vertically on the side walls of the room or horizontally on the
ceiling above the center isle. An alternative is to mount the lights on the underside of each tier of
shelf or tray, at least 18 inches above the cropping surface. To eliminate the heat and consequent
drying action caused by the fixture ballasts, these can be removed and placed outside the room.

The best type of light tube is one which most closely resembles natural outdoor light: i.e. one
that has at least 140 microwatts per 10 nanometer per lumen of blue spectra (440-495 nm). In
contrast, warm-white fluorescent light has only 40-50 microwatts/nm/lum. and cool-white has
100-110 microwatts/nm/lum. Commercial lights meeting the photo-requirements of species
mentioned in this book are the “Daylite 65” kind manufactured by the Durotest Corporation and
having a “color temperature” of 6500 ° K and the “Vita-Lite” fluorescent at 5500 ° K. These color
temperatures provide the proper amount of blue light for promoting primordia formation in
Pleurotus ostreatus, Psilocybe and in other photosensitive species.

Environmental Monito^/c5|zquipment

Few organisms are as sensitive to fluctuations in the environment as mushrooms. A matter of a
few degrees in temperature or humidity can dramatically influence the progression of fruiting and af-
fect overall yields. To adequately monitor the growing environment, quality equipment is essential
for accurate readings. This equipment should include maximum-minimum thermometers to gauge
temperature fluctuations and a hygrometer or a sling psychrometer for measuring humidity. Hygro-
meters should be periodically calibrated with a sling psychrometer to insure accuracy. Thermome-
ters also should be checked as there are occasional irregularities. Other more advanced, expensive
but not absolutely essential equipment helpful to mushroom growers include: C0 2 detectors;
moisture meters; anemometers; and light measuring devices.

78/The Mushroom Cultivator

T he purpose of composting is to prepare a nutritious medium of such characteristics that the
growth of mushroom mycelium is promoted to the practical exclusion of competitor organ-
isms. Specifically this means:

1 . To create a physically and chemically homogeneous substrate.

2. To create a selective substrate, one in which the mushroom mycelium thrives better than
competitor microorganisms.

3. To concentrate nutrients for use by the mushroom plant and to exhaust nutrients favored by
competitors.

4. To remove the heat generating capabilities of the substrate.

Mushroom mycelium grows on a wide variety of plant matter and animal manures. These
materials occur naturally in various combinations and in varying stages of decomposition. Physically
and chemically they are a heterogeneous mixture containing a wide variety of insects, microorgan-
isms and nematodes. Many of these organisms directly compete with the mushroom mycelium for
the available nutrients and inhibit its growth. By composting, nutrients favored by competitors grad-
ually diminish while nutrients available to the mushroom mycelium are accumulated. With time, the
substrate becomes specific for the growth of mushrooms.

The composting process is divided into two stages, commonly called Phase I and Phase II.
Each stage is designed to accomplish specific ends, these being:

Phase I: Termed outdoor composting, this stage involves the mixing and primary decomposi-
tion of the raw materials.

Phase II: Carried out indoors in specially designed rooms, the compost is pasteurized and
conditioned within strict temperature zones.

Basic Raw Materials

The basic raw material used for composting is cereal straw from wheat, rye, oat, barley and rye
grass. Of these, wheat straw is preferred due to its more resilient nature. This characteristic helps
provide structure to the compost. Other straw types, oat and barley in particular, tend to flatten out
and waterlog, leading to anaerobic conditions within a compost pile. Rye grass straw is more resis-
tant to decomposition, taking longer to compost. Given these factors and with proper management,
all straw types can be used successfully.

Straw provides a compost with carbohydrates, the basic food stuffs of mushroom nutrition.
Wheat straw is 36% cellulose, 25% pentosan and 16% lignin. Cellulose and pentosan are carbo-
hydrates which upon break down yield simple sugars. These sugars supply the energy for microbial
growth. Lignin, a highly resistant material also found in the heartwood of trees, is changed during
composting to a “Nitrogen-rich-lignin-humus-complex”, a source of protein. In essence, straw is a
material with the structural and chemical properties ideal for making a mushroom compost.

When cereal straw is gathered from horse stables, it is called “horse manure’ . Although culti-
vators call it by this name, the material is actually 90% straw and 10% manure. This “horse ma-
nure” includes the droppings, urine and straw that has been bedding material. The quality of this

Compost Preparation/79

material depends on the proportions of urine and droppings present, the essential elements nitro-
gen, phosphorous and potassium being contained therein. The reason horse manure is favored for
making compost is the fact that fully 30-40% of the droppings are comprised of living microorgan-
isms. These microorganisms accelerate the composting process, thereby giving horse manure a
decided advantage over other raw materials.

Horse manure used by commercial mushroom farms generally comes from race tracks. The
bedding straw is changed frequently, producing a material that is light in urine and droppings. On
the other hand, boarding stables change the bedding less, generating a heavier material. If sawdust
or shavings are used in place of straw for bedding, the material should be regarded as a supplement
and not as a basic starting ingredient.

When horse manure is used as the basic starting ingredient, the compost is considered a
“horse manure compost” whereas “synthetic compost” refers to a compost using no horse ma-
nure. Straw, sometimes mixed with hay, is the base ingredient in synthetic composts. Because straw
is low in potassium and phosphorus, these elements must be provided by supplementation and for
this reason chicken manure is the standard additive for synthetic composts. No composts are made
exclusively of hay because of its high cost and small fiber. In fact, mushroom growers have tradition-
ally used waste products because they are both cheap and readily available.

By themselves horse manure or straw are insufficient for producing a nutritious compost. Nor
do they decompose rapidly. They must be fortified by specific materials called supplements. In
order to determine how much supplementation is necessary for a given amount of horse manure or
straw based synthetic, a special formula is used. This formula insures the correct proportion of initial
ingredients, which largely determines the course of the composting process. The formula is based
on the total nitrogen present in each ingredient as determined by the Kjeldhal method. By using this
formula and certain composting principles, the carbomnitrogen ratio for optimum microbial decom-
positions is assured. In turn, maximum nutritional value will be achieved.

Composting is a process of microbial decomposition. The microbes are already present in
large numbers in the compost ingredients and need only the addition of water to become active. To
stimulate microbial activity and enhance their growth, nutrient supplements are added to the bulk
starting materials. These supplements are designed to provide protein (nitrogen) and carbohydrates
to feed the ever increasing microbial populations. Microbes can use almost any nitrogen source as
long as sufficient carbohydrates are readily available to supply energy for the nitrogen utilization. Be-
cause of the tough nature of cellulose, the carbohydrates in straw are not initially usable and must
come from another source. A balanced supplement is therefore highly desirable. It should contain
not only nitrogen but also sufficient organic matter to supply these essential carbohydrates. For this
reason certain manures and animal feed meals are widely used for composting.

The following is a list of possible compost ingredients or supplements, grouped according to
nitrogen content. Their use by commercial growers is largely determined by availability and cost.
This list is not all inclusive and similar materials can be substituted. (See Appendix).

80/The Mushroom Cultivator

Group I: High nitrogen, no organic matter

Ammonium sulfate— 21% N
Ammonium nitrate — 26% N
Urea — 46% N

Maximum rate— 25 Ibs/dry ton of starting materials

These are inorganic compounds that supply a rapid burst of ammonia. They are frequently
used for initial straw softening in synthetic composts. When used, care should be taken that they are
applied evenly. If ammonium sulfate is used, calcium carbonate must also be added at a rate of 3
parts CaC0 3 to 1 , to neutralize sulfuric acid groups. These supplements are not recommended for
horse manure composts.

Group II: 10-14% N

Blood Meal — 13.5% N
Fish Meal — 10.5% N

These materials consist mainly of proteins but because of their high cost are rarely used.

Group III: 3-7% N

Malt sprouts— 4% N

Brewers’ grains— 3-5% N

Cottonseed meal — 6.5% N

Peanut meal— 6.5% N

Chicken manure— 3-6% N

This group contains the materials most widely used by commercial growers and is character-
ized by a favorable carbomnitrogen balance. Dried chicken manure from broilers mixed with saw-
dust is commonly used and easy to handle.

Group IV: Low nitrogen, high carbohydrate

Grape pomace — 1.5% N
Sugar beet pulp — 1.5% N
Potato pulp — 1 % N
Apple pomace — 0.7% N
Molasses - 0.5% N
Cottonseed hulls — 1% N

These materials are excellent temperature boosters and for this reason are a recommended
additive to all composts. They can be added to any compost formula at a rate of 250 lbs per dry to
of ingredients. Cottonseed hulls are an excellent structural additive.

Group V: Animal manures

Cow manure — 0.5 % N
Pig manure — 0. 3-0.8% N

These manures are rarely used for composting, except in areas without horses or chickens.
They have been used with success and should be considered supplements to a synthetic blend.

Compost Preparation/81

Group VI: Hay

Alfalfa-2.0-2. 5% N
Clover— 2% N

Hay is useful for boosting initial temperatures in synthetic composts. Hay contains substantial
quantities of carbohydrates which help build the microbial population. Yet another advantage is the
relatively high nitrogen content in alfalfa and clover. Use at a rate of 20% of the basic starting mate-
rial (dry weight).

Group VII: Minerals

Gypsum — Calcium sulfate

Gypsum is an essential element for all composts. Its action, largely chemical in nature, facili-
tates proper composting. Its effects are:

1 . To improve the physical structure of the compost by causing aggregation of colloidal parti-
cles. This produces a more granular, open structure which results in larger air spaces and
improved aeration.

2. To increase the water holding capacity, while decreasing the danger of over-wetting. Loose
water is bound to the straw by colloidal particles.

3. To counteract harmfully high concentrations of the elements K, Mg, P and Na should they
occur, thereby preventing a greasy condition in the compost.

4. To supply the calcium necessary for mushroom metabolism.

Gypsum should be added at a rate of 50- 1 00 lbs per dry ton of ingredients. When supplement-
ing with chicken manure, it is advisable to use the high rate.

Limestone flour — Calcium carbonate

Limestone is used when one or more supplements are very acidic and need to be buffered. A
good example of this is grape pomace, which has a pH of 4. Because it is added in large quantities,
grape pomace could affect the composting process which normally occurs under alkaline condi-
tions.

Group VIII: Starting materials

Horse manure — 0.9-1 .2% IN
Straw, all types— 0.5-0. 7% IN

Compost Formulas

The following formulas for high yield compost are commercially proven. If an ingredient is not
available locally, substitute one that is. The aim of the formula is to achieve a nitrogen content of
1 .5-1 .7% at the initial make-up of the compost pile.

In order for these formulas to be effective, the moisture content and nitrogen content must be
correct. Moisture level is determined by weighing 1 00 grams of the material, drying it in an oven at
200 ° F. for several hours, and then reweighing it. The difference is the percent moisture. Be sure
the sample is representative. The nitrogen content (protein divided by 6.25) is always listed with

82/The Mushroom Cultivator

commercial materials because they are priced according to percentage of protein. On the other
hand, barnyard materials vary considerably with age. The more a material breaks down, the more
nitrogen it loses. Most compost supplements are purchased dry and added dry, helping even distri-
bution as well as enabling easy storage. It is also important that the raw materials used for compost-
ing be as fresh as possible. This insures maximum utilization of their properties. Baled straw stored
for a year and kept dry is fine. If the straw has gotten wet, moldy or otherwise started to decompose,
it should not be used.

Formula I

Ingredient

Wet u/t.

%h 2 o

Dry wt. %N

lbs.

Horse manure

2,000

1,000 1.0

Cottonseed meal

117 6.5

Gypsum

—

50 -

1,167

(18) + (1,1 67) = 1 .54% N

This formula makes approximately 2800 pounds of compost

at a 70% moisture content.

Formula II

Ingredient

Wet u/t.

%h 2 o

Dry u/t. %N

Ibs.N

Wheat straw

2,000

1,800 0.5

Chicken manure

2,000

1,600 3.00

Gypsum

125

— -

125

3,525

(57) (3,525) = 1.62%N

This formula make approximately 7,000 pounds of compost at a 71 % moisture content.

Although 7,000 pounds of compost seems like a large quantity, at a fill level of 20 pounds per
sq. ft., this will fill only 350 sq. ft. of beds or trays. Keep in mind that during the composting process
there is a gradual reduction in the the total volume of raw materials. Fully 20-30% of the dry matter
is consumed during Phase I and another 10-1 5% during Phase II. In total, approximately 40%
of the dry matter is reduced by microbial and chemical processes. This loss of potential nu-
trients can not be avoided and demonstrates the importance of composting no longer than neces-
sary.

Ammonia

The production of ammonia is essential to the composting process. Just as the carbohydrates
must be in a form that microbes can utilize, so must the nitrogen.

1 . Ammonia supplies nitrogen for microbial use.

2. Ammonia is produced by microbes acting upon the protein contained in the supplements.

With the energy supplied by readily available carbohydrates, microbes use the ammonia to

Compost Preparation/83

form body tissues. A microbial succession of generations is established, with each new generation
decomposing the remains of the previous one. Microbial action also fixes a certain amount of the
ammonia, forming the “nitrogen-rich-lignin-humus-complex”. Unused ammonia volatilizes into the
atmosphere. The smell of ammonia should be evident throughout Phase I. reaching a peak at fill-
ing.

CarbomNitrogen Ratio

The importance of a carbonmitrogen balance cannot be underestimated. A well balanced com-
post holds an optimum nutritional level for microbial growth. An imbalance slows and impedes this
growth. It is the compost formula that enables the grower to achieve the correct C:N balance. Be-
cause organic matter is reduced during composting, the C:N ratio gradually decreases. Approxi-
mate values are: 30: 1 at make-up; 20:1 at filling; and 17:1 at spawning.

1 . Over-supplementation with nitrogen results in prolonged ammonia release.

2. Over-supplementation with carbohydrates results in residual carbon compounds.

Prolonged ammonia release from an over-supplemented compost necessitates longer com-
posting times. If composting continues too long, the physical structure and nutritional qualities are
negatively affected. If the ammonia persists, the compost becomes unsuitable for mycelial growth.

Readily available carbohydrates which are not consumed by the microbes during composting
can become food for competitors. It is therefore important that these compounds are no longer
present when composting is finished.

Water and Air

Water is the most important component in the composting process. To a large degree water
governs the level of microbial activity, in turn, this activity determines the amount of heat generated
within the compost pile because the microorganisms can only take up nutrients in solution. Not
only do the microorganisms need water to thrive, but they also need oxygen. Years of practice and
research have established a basic relationship between the amount of water added and the aeration
of the compost. An inverse relationship exists between the amount of water and the
amount of oxygen in a compost pile.

1. Too much water = too little air
Moisture content 75% or above.

2. Too little water = too much air
Moisture content 67% or below

Overwetting a compost causes the air spaces to fill with water. Oxygen is unable to penetrate,
causing an anaerobic condition. In contrast, insufficient water results in a compost that is too airy.
Beneficial high temperatures are never reached because the heat generated is quickly convected
away.

84/The Mushroom Cultivator

Pre-wetting

As long as the composting ingredients remain dry, the microorganisms lie dormant and com-
posting does not take place. The first step in the composting process is the initial watering of the
starting materials. The purpose of this pre-composting or pre-wetting is to activate the microbes.

Compost microorganisms can be divided into two classes according to their oxygen require-
ments. Those needing oxygen to live and grow are called Aerobes while those living in the ab-
sence of oxygen are called Anaerobes. Each class has well defined characteristics.

t. Aerobes decompose organic matter rapidly and completely with a corresponding produc-
tion of C0 2 , water and heat. This heat generation is called Thermogenesis.

2. Anaerobes partially decompose organic matter, producing not only C0 2 and water, but
also certain organic acids and several types of gases such as hydrogen sulfide and methane.
Anaerobes generate less heat than aerobes.

Examination of anaerobic areas of the compost reveals a yellowish, under-composted material
that smells like rotten eggs. These areas in a compost pile are noticeably cooler and generally water-
logged. Anaerobic compost is unsuitable for mushroom growth.

Since neither fresh horse manure nor straw based synthetics have the correct moisture content,
water must be added to these materials. The recommended levels for optimum composting are:

Horse manure: 69-71% Synthetic: 71-73%

vi'T

Figure 83 Pre-wetted raw materials in a windrow.

Compost Preparation/85

E32

Once activated, the microbes begin to attack the straw and decompose the waxy film which encases
the straw fibers. Until this film is degraded, water will not penetrate the straw and its nutrients will re-
main unavailable. As the process progresses, the fibers become increasingly receptive to water,
which rather than being free or on the surface, penetrates and is absorbed into the straw.

There are many methods for pre-wetting. These include: dipping or dunking the material into a
tank of water; spraying it with a hose; or spreading it out in a flat pile 2-3 feet high and running a
sprinkler over it. Regardless of the method used, the result should be the same— a homogeneous
evenly wetted pile.

Horse manure needs less time for pre-wetting due to the nature of the bedding straw. This straw
has been trampled upon, opening the straw fiber and damaging the waxy film. The urine and drop-
pings have also begun to soften it. This is not the case with a synthetic compost in which the baled
straw is still fresh and tough. To stimulate microbial action in synthetics, some supplements are
added at pre-wetting. Suitable supplements include any from group 1 , 4, 5 or chicken manure.

The length of time needed for pre-wetting varies according to the condition of the starting mate-
rials. Generally 3 days for horse manure and 5-12 days for a synthetic compost is sufficient. The
pre-wetting time for a synthetic compost can be shortened if the straw is mechanically chopped, but
care should be taken that the fibers do not become too short.

The wetted materials are then piled in a large rounded heap called a windrow. During this peri-
od the windrow can be turned and re-wetted as needed, usually 1-3 times.

Building the Pile

Building the compost pile is called stacking, ricking or “make-up”. At this time the pre-wetted
starting materials and the nitrogenous supplements are evenly mixed, watered and assembled into a
pile. The size, shape and specific physical properties of this pile are very important for optimum
composting. These are:

1 . Pile dimensions should be 5-6 feet wide by 4-6 feet high. The shape should be rectangular
or square.

2. The side of the pile should be vertical and compressed from the outside by 3-6 inches. The
internal section should be less dense than the outer section.

3. The pile is such that any further increase in size would result in an anaerobic core.

Throughout the composting process, the size of the pile varies depending on the physical con-
dition of the straw, which provides the pile’s basic structure. The structure of the compost refers to
the physical interaction of raw materials, especially the straw fibers. As the straw degrades and the fi-
bers flatten out, the structure becomes more dense and the airflow is restricted, The pile becomes
more compact and its size is reduced accordingly. Initially the fresh straw allows for generous air
penetration which convects away heat and slows microbial action. To counteract this heat loss, the
pile should be of maximum size and optimum moisture content at make-up.

Figure 85 illustrates air penetration of a compost pile. Air enters the pile from the sides. As the

86/The Mushroom Cultivator

Compost Preparation/87

oxygen is used by microorganisms, heat is set free and the air temperature rises. The warm air cur-
rent created rises to the top of the pile. This is called the chimney effect. The factors that affect the
rate of internal air flow are pile size and structure, moisture content and the differential between am-
bient air and internal pile temperatures.

ZuS

riposting.

Turning

A well built compost pile runs out of oxygen in 48 to 96 hours and then enters an anaerobic
state. To prevent this, the pile should be disassembled and then reassembled. The purposes of this
turning procedure are:

1 . To aerate the pile, preventing anaerobi^,

2. To add water lost through evaporation

3. To mix in supplements as required.

4. To fully mix the compost, preventing uneven decomposition.

As a consequence of microbial decomposition, the compost pile begins to shrink and becomes
more compact. Coupled with loose water gravitating downward and water generated by microbes in
the inner active areas, this compaction closes the air spaces and stifles aerobic action, particularly in
the core at the bottom center. Through the use of a long stemmed thermometer reaching to the
center of the pile, the time of oxygen depletion can be monitored by watching temperature. When
the temperature begins to drop, indicating a slowing of microbial action, it is time to turn the com-
post.

88/The Mushroom Cultivator

D-'-n V't-'r M m, I BAM M— ^ 1 ' ~ —— 1 ■

In the early stages the temperature stratification in the pile is quite pronounced. Outer areas are
cool and dry from the air flowing inward and the accompanying evaporation. These outer areas are
watered during turning and moved to the center of the newly built pile and the center areas are relo-
cated to the outside. Being aware of the varied rate of decomposition in a stratified pile and compen-
sating during turning maintains the important homogeneous character of the pile.

Supplements deleted at make-up should be added during the turning cycle. Gypsum is normal-
ly added at the second turn. Adding gypsum any earlier is believed to depress ammonia production.
Until some decomposition has occurred, the beneficial action of gypsum will not be realized. As
with other supplements, gypsum is mixed in as evenly as possible.

Temperature

Environmental conditions in the compost are specifically designed to facilitate growth of benefi-
cial aerobic microorganisms. Given the proper balance of raw materials, air and water, a continuous
succession of microbial populations produces temperatures up to 180°F. These microbes can be
divided into two groups according to their temperature requirements. Mesophiles are active under
90°F. and thermophiles are active from 90-160 °F. The action of these microbial groups during
the composting process is summarized in the following paragraphs.

During pre-composting mesophilic bacteria and fungi, utilizing available carbohydrates, attack

Figure 87 Standard temperature zonation in a compost pile.

Compost Preparation/ 89

the nitrogenous compounds thereby releasing ammonia. This ammonia is then utilized by succes-
sive microbial populations and the temperature rises.

After make-up, the mesophiles remain in the cool outer zones while the thermophilic fungi, ac-
tinomycetes and bacteria dominate the inside of the pile. The actinomycetes are clearly visible as
whitish flecks forming a distinct ring around the hot center. Bacteria dominate this center area and
continue to decompose the nitrogenous supplements, liberating more ammonia. At this point the
carbohydrates in the straw are ready for microbial use.

At temperatures over 1 50 °F., microbial action slows and chemical processes begin. Between
1 50-165 °F. microbial and chemical actions occur simultaneously. From 1 65-1 80 °F. decomposi-
tion is mainly due to the chemical reactions of humification and caramellization, the latter taking
place under conditions of high temperature, high pH (8.5) and in the presence of ammonia and
oxygen. Many of the dark compounds produced during composting are believed to result from
these chemical reactions. Decomposition proceeds rapidly at these high temperatures, and if they
can be maintained throughout the process, composting time will be greatly reduced.

Figure 87 shows the temperature zonation commonly found in a compost pile. Studies by Dr.
E.B. Lambert in the 1 930’s showed that compost taken from zone 2 produced the highest yielding
crops. Based on this research, growers always subject their compost to zone 2 conditions prior to
spawning. This normally occurs during Phase II in specially designed rooms. However, if a Phase 11
room can not be built, zone 2 conditions can be achieved by an alternate method known as Long
Composting, developed by C. Riber Rasmussen of Denmark.

Long Composting

Long composting is designed to carry out the complete composting process outdoors (exclud-
ing pasteurization). The method is characterized by the avoidance of high temperature chemical de-
composition and a reliance on purely microbial action. Specifically this procedure is designed to
promote the growth of actinomycetes and rid the compost of all ammonia by the time of filling. The
temperature zonation desired in this method is illustrated in Figure 88. An outline of the Long Com-
posting procedure follows.

DAY LONG COMPOSTING PROCEDURE

-10 For synthetic composts: Break the straw bales and water them thoroughly. Mix in
group 1 ,4 or 5 supplements or chicken manure. Windrow. Start at day -5 if straw is
short or has been chopped.

-5 For synthetic composts: Turn and add more water. Break up any concentrations of
supplements. Windrow.

-2 For horse manure or synthetic composts: Thoroughly wet and mix all raw materials
and supplements (except gypsum). Windrow.

0 Make up the pile. Dimensions should be 6 feet wide and 4 feet high. The vertical sides

90/The Mushroom Cultivator

should be tightly compressed with the middle of the pile remaining loose. Use the pile
formers to make the stack and stomp the sides from the top to achieve ample compres-
sion. Water dry areas.

6 First turn: Water as needed. Move the center anaerobic zone to the outside of the new
pile and the outside zone to the center. Keep the pile height and length constant by re-
ducing the width as decomposition proceeds.

10-12 Second turn: Add the gypsum and water as needed. Distribute the zone of actinomy-
cetes evenly throughout.

13-15 Third turn: The actinomycete zone should be evident throughout. Strong actinomy-
cete growth may cause excessive drying, so be sure to check moisture content and water
as needed. The smell of ammonia should be slight. Build the new pile only 24 inches
high and 4-5 feet wide. Distribute the actinomycetes evenly throughout.

15-17 Fourth turn: The compost should now appear dark brown and well flecked with actino-
mycetes. All traces of ammonia should be gone. Moisture content should be approxi-
mately 67-70% and the pH 7. 0-7. 5. If this is not the case, continue the process turning
at 2 day intervals until this condition is reached. The pile height may vary between
1 6-24 inches and is designed solely to promote optimum conditions for the growth of
the actinomycetes— temperatures of 120-1 35 °F.

Once finished, this compost is normally pasteurized at 1 35 °F. for four hours. If pasteurization
is impossible, discard the cool outer shell and utilize the areas showing strong actinomycete activity.
Although these areas will not be free from all pests and competitors, they should provide a reas-
onably productive substrate. The aspect and characteristics of a properly prepared Long Compost
should conform to the guidelines for compost after Phase II. (See Aspect of the Finished Compost
on page 105 and Color Plate 8).

Short Composting

Commercial Agaricus growers uniformly base their composting procedures on the methodolo-
gy developed by Dr. James Sinden, who called his technique “Short Composting” in reference to
the short period of time involved. Dr. Sinden’s process is centered around the fast acting chemical
reactions occurring in zone 3. Besides the shorter preparation time, this process also results in a
greater preservation of dry matter, thus retaining valuable nutrients. Figure 89 illustrates the zona-
tion during short composting.

Without commercial composting equipment, approximating the temperature conditions of
Short Composting is very difficult. However, it does provide a model for optimum composting and
can be approached by adhering to the basic principles discussed in this chapter. The Short Com-
posting procedure is outlined below.

Compost Preparation/91

DAY SHORT COMPOSTING PROCEDURE; Formula 1

-1 For horse manure: Wet the starting materials thoroughly. Windrow.

0 Make up the pile. Add all supplements except gypsum. Mix and water thoroughly. Pile

should be 6 feet wide by 5-6 feet high. The sides should be vertical and compressed
tightly.

2,3 First turn: Add gypsum and water as needed. Keep the pile height constant and vary
the width only in relation to the amount of anaerobic material.

5 Second turn: Add water as needed.

7 Third turn: Add water as needed. Compost should be ready to fill.

The procedures for making a synthetic compost by the short composting method are outlined

below, with minor modifications for the home cultivator. Note the longer period of pre-composting

to condition the straw.

DAY SYNTHETIC COMPOSTING PROCEDURE: Formula 2

- 1 0 Break straw bales and wet thoroughly. Windrow or spread out in a low flat pile, 2-3 feet

high. Water daily.

-7 Mix the chicken manure together with the straw, wetting both well. Avoid water run-off.

Windrow.

-3 Re-mix the windrow, adding water as necessary. Start here if chopped straw is used-wet

the straw and chicken manure. Mix well and windrow.

0 Make up the pile. Dimensions should be 6 feet wide by 5 feet high. Add as much

water as possible without run-off. Use pile formers to insure vertical sides and stomp

Figure 88 Temperature zonation during Figure 89 Temperature zonation during
Long Composting. Short Composting.

92/The Mushroom Cultivator

Figure 90 Commercial compost turning machine.

down the sides from the top to achieve adequate compression. The pile should be tight
and compact.

4 First turn: Add the gypsum. Water as needed. Keep the pile dimensions constant, vary-

ing the width as indicated by the amount of anaerobic material in the center. Maintain
pile compaction.

7 Second turn: Water as needed and redistribute outer and inner areas. Redistribution

should occur during each turn to keep the material in an even state of decomposition.
10 Third turn: Mix well and add water as needed. Reduce width to 5 feet. Fill if ready.

13 Fill if ready, or continue composting, fuming at two day intervals.

Composting Tools

Since commercial growers work with many tons of compost, a bucket loader is essential. They
also use a specially designed machine for turning the piles. This compost turner can travel through
a 200 foot pile in a little over one hour, mixing in supplements and adding water. Small scale culti-
vators can make compost without these machines. The following is a list of tools and facilities that
are basic to compost preparation.

1 . A cement floor. Not absolutely necessary but highly desirable, a cement floor is easy to
work on, prevents migration of water to the earth and prevents soil and unwanted soil or-
ganisms from contaminating the pile. Water leaching from the pile, a good indicator of
compost moistures, is quite evident on a cement floor. If a cement floor is not available, a
sheet of heavy plastic can be used.

2. Bobcat or small tractor loader with 3 A- 1 yard bucket with fork. If producing large
amounts of compost, one of these machines saves time and labor. Not only do they make

Compost Preparation/93

Figure 91 Pile formers in use.

pre-wetting, supplementing and pile building easier, they can be used to turn the pile.

3. Pile formers. These are constructed from 2 x 4’s and plywood or planks to the dimen-
sions desired for the compost pile One for each side is necessary. Standard size would be
4-5 feet high by 8 feet long. An alternative to pile formers is a three sided bin.

4. Long handled pitchfork with 4 or 5 prongs. The basic tool in a compost yard, all com-
post piles were turned with pitchforks before the advent of compost turners and bucket
loaders.

5. Flat bladed shovel. Used for handling supplements.

6. Hose with spray nozzle, or sprinkler.

7. Thermometers. Although pile temperatures can be guaged by touch, a long stemmed
thermometer gives accurate readings.

Characteristics of the Compost at Filling

The composting materials undergo very distinct changes during Phase I. A judgment as to the
suitability of the compost for filling is based on color, texture and odor. Gradual darkening of the
straw and the pronounced scent of ammonia are the most obvious features. These and other char-
acteristics provide important guidelines for judging the right time for filling the compost. (Note:
these guideslines do not apply for a compost prepared by the Long Composting methods.)

The compost is ready for filling if:

1 . Compost is uniformly deep brown.

2. Straw is still long and fibrous, but can be sheared with some resistance.

3. When the compost is firmly squeezed, liquid appears between the fingers.

94/The Mushroom Cultivator

Figure 92, 93 Compost at
filling can be sheared with
moderate resisance.

Figure 94 Compost at fill-
ing should release some
moisture when firmly
squeezed.

mf ' v

/ 1 Pm

. m

Compost Preparation/95

4. Compost has a strong smell of ammonia, pH of 8. 0-8. 5.

5. Compost is lightly flecked with whitish colonies of actinomycetes.

6. Kjeldahl nitrogen is 1 .5% for horse manure and 1 .7% for synthetic composts.

Supplementation at Filling

The key to a successful Phase II, whether in trays, shelves or a bulk room, lies in the heat gen-
erating capabilities of the completed Phase I compost. To this end the compost should be biologi-
cally “active,” a term that describes a compost with sufficient food reserves to sustain a high level of
microbial activity. Whereas the Sinden Short Compost is a model of a vitally active compost, the
Rasmussen Long Compost is considered biologically “dead” because these food reserves have
been deliberately exhausted during Phase I. In this same sense, a compost having completed the
Phase II is also considered a dead compost.

A method that insures a high level of microbial activity during the Phase II is supplementation
with highly soluble carbohydrates during Phase I or with vegetable oils (fats) at filling. The purpose
of these supplements is to provide readily available nutrients which stimulate the growth of the mi-
crobial populations. The effect of carbohydrates or oil supplementation on the Phase II is:

1 . Accelerated thermogenesis— The nutrients provided by the supplements act as a “super-
charger” for the microbial populations. Consequently their increased activity generates
more heat. Specifically, supplementation with vegetable oil (cottonseed oil) increased popu-
lations of actinomycetes and thermophilic fungi (Schisler and Patton, 1 970) while soluble
carbohydrates (molasses) enhanced bacterial populations (Hayes and Randle, 1968).

2. Better compost ventilation— Heightened thermogenesis within the compost requires lower
air temperatures within the Phase II room. The greater the compost to air temperature dif-
ferential, the better the air movement through the compost. In this respect a dead compost
requires a high room temperature and is difficult to condition because of its low microbial
activity.

3. Rapid reduction of free ammonia— The increased ventilation and microbial activity give rise
to a rapid fixation of ammonia. As a result, the Phase II period is reduced by as much as
three days. The advantage of this reduced time period is that dry matter and hence nutrients
for mushroom growth are conserved.

4. Reduced spawn running period— Oil supplemented composts show increased mycelial ac-
tivity and therefore higher temperatures during the spawn running period. As a result the
colonization period is shortened by three to five days.

5. Increased yields— Yield increases of 0.4-0. 5 lbs/ft 2 are common for Agaricus growers us-
ing vegetable oil at filling. Similar increases are reported for molasses.

Compost supplementation with soluble carbohydrates is an effective way to prepare an active com-
post. These materials are listed earlier in the chapter as Group IV supplements. They are added to a
synthetic compost during pre-composting (50%) and at third turn (50%) and to a horse manure
compost at make-up and at third turn. Molasses is added at make-up at a rate of 1 0 ml per pound of

96/The Mushroom C ultivator

compost wet weight and is diluted 1 :2 with water for easy application. Vegetable oil is sprayed onto
the compost the day of fill at a rate of 1 0 ml per pound of compost wet weight. Even application is
important to avoid creating hot spots.

Compost supplementation with soluble carbohydrates or vegetable oils is highly recom-
mended, especially for those planning a Phase 11 without steam or with only limited supplemental
heating. Hence, this type of supplementation is particularly appropriate for the home cultivator.

While Phase I is a combination of biological and chemical processes, Phase II is purely biologi-
cal. In fact, Phase II can be considered a process of microbial husbandry. By bringing the compost
indoors into specially designed rooms, the environmental factors of temperature, humidity and fresh
air can be controlled to such a degree that conditions for growth of select microbial groups can be
maximized. These thermophilic and thermotolerant groups and their temperature ranges are:

Bacteria: 1 00-1 70 °F. Different species of bacteria are active throughout this range so an op-
timum can not be given. At temperatures above 1 30°F. bacteria dominate and are responsible for
the ammonification that occurs at these temperatures. The most common bacteria found by resear-
chers are Pseudomonas species.

Actinomycetes: 1 1 5-140°F. with an optimum temperature range of 125-132°F . The most
common species are found in the genera Streptomyces and Thermomonospora. Work done by
Stanek (1971) has shown that actinomycetes and bacteria are mutually stimulatory, resulting in
greater efficiency when working together.

Fungi: 110-1 30 °F. with an optimum temperature of 118-122°F. Common genera are
Humicola and Torula. Recent research indicates that these fungi are the most efficient de-ammoni-
fiers, which has led to a more general use of their temperature range for Phase 11 conditioning.

The basic function of these microorganisms is to utilize and thereby exhaust the readily availa-
ble carbohydrates and the free ammonia. Ammonia in particular must be completely removed be-

Figure 95 Temperature vs.
ammonia utilization by
microbial populations.

(After Ross, 1978)

Compost Preparation/97

cause of its inhibitory effect on the growth of mushroom mycelium. The result of this microbial ac-
tion is a build-up of cell substance or “biomass” which contains vitamins, fats and proteins. What
the mushroom mycelium uses for a large portion of its nutrition then, is the concentrated bodies
forming the microbial biomass. This biomass constitutes part of the brown layer coating the partially
decomposed straw fibers.

Many growers consider Phase II to be the most important stage in the growing cycle and rightly
so. An improperly prepared substrate yields few if any mushrooms. It is critical, therefore, that the
environmental conditions required during Phase II be carefully maintained. Phase 1 1 can be sepa-
rated into two distinct parts, each serving a specific function. These are:

1 . PASTEURIZATION: The air and compost temperature are held at 1 35-140 °F. for 2-6

hours. The purpose of pasteurization is to kill or neutralize all harmful organisms in the
compost, compost container and the room. These are mainly nematodes, eggs and larvae
of flies, mites, harmful fungi and their spores. The length of time needed generally depends
on the depth of fill. Deeper compost layers require more time than shallow ones. In gener-
al, two hours at 140°F. is sufficient. Compost temperatures above 140°F. must be avoid-
ed because they inactivate fungi and actinomycetes while at the same time stimulating the
ammonifying bacteria. If temperatures do go above 140°F,, be sure there is a generous
supply of fresh air. , 28

2. CONDITIONING: The compost temperature is held at 1 1 jp Once the pasteuriza-
tion is completed, the compost temperature should be lowerW^|l|ally over 24 hours to
the temperature zone favored by actinomycetes and fungi. Th^exaHhemperature varies ac-
cording to the depth of fill. At depths up to 8 inches, 1 22 °F. as measured in the center of
the compost is most frequently used. At depths over 8 inches, temperature stratification
becomes more pronounced, making a higher core temperature of 1 28 °F. advantageous. A
common procedure is to bring the compost temperature down in steps, dropping the core
temperature 2 °per day, from 1 30 ° to 122 °F. This temperature is then held until all traces
of ammonia are gone.

Basic Air Requirements

Phase II is purely a process of aerobic fermentation and as such a constant supply of fresh air is
essential. To insure this supply, a minimum fresh air setting is established on the air intake damper.
A standard minimum setting is 8-10% of the intake opening. The oxygen level can be checked in a
practical manner by lighting a match in the Phase II room. If a flame can be maintained, the oxygen
level is sufficient. Lack of oxygen stimulates the growth of Chaetomium, the Olive Green Mold,
which will spoil the compost. (See Chapter XIII).

Compost temperatures follow the air temperature of the room. Fresh air not only supplies oxy-
gen, but is also used to keep the compost within the correct temperature zone. To drop the com-
post temperature, more fresh air is introduced and vice versa. Oversupply of fresh air is only a prob-
lem if it leads to rapid cooling of the compost. In this regard, changes in the fresh air setting should

98/The Mushroom Cultivator

be slow and deliberate. Only when the compost threatens to overheat should maximum fresh air be
introduced. This is particularly common directly after pasteurization.

Peak microbial activity normally occurs 24-48 hours after pasteurization. As Phase II pro-
gresses and the food supply diminishes, this activity begins to slow. Compost temperatures should
begin to drop on their own. As they drop, the fresh air supply should be decreased, thus slowly rais-
ing the air temperature as the compost reaches the required temperature zones. If the fresh air
minimum is reached and the compost temperatures are still dropping, a supplemental heat source
must be installed.

Phase II Room Design

The Phase II room can be a special room set aside solely for this purpose (the norm on fray
farms) or it can be in the same room where cropping occurs. Design features are critical for its suc-
cess and should be strictly adhered to. These features are:

1. Adequate insulation: Insulate to a R value of 19 for walls and a minimum of 30 for the ceil-
ing. A vapor barrier is needed to protect the insulation. (A layer of polyethylene is cheap and
effective.)

2. The room must be functionally airtight. The door should form a tight seal. Any cracks or
openings allow the passage of flies.

3. The ventilation system uses a backward-curved centrifugal fan driven by pulleys and belts,
and whose speed can be varied. The fan should be capable of moving air at 1 cubic foot per
minute (CFM) per square foot of compost surface area. A perforated polythene duct runs the
length of the room and directs the air either straight down the center aisle or across the ceil-
ing to the side walls. High velocity airflow is necessary to maintain even temperatures
throughout as well as to keep the room under positive pressure.

4. A fresh air vent is located before the fan. This damper also regulates recirculated air. (See
Fig. 73).

5. Filters are placed before the fresh air inlet. These filters are important as protection against
flies, dust and spores. High efficiency spore filters are commonly used for the incoming fresh
air. A pre-filter placed upstream of the main filter will increase its life. Recirculated air should
never be filtered during Phase II because of its high moisture content.

6. At the opposite end of the room from the fresh air vent are exhaust louvers operating on air
pressure. This exhaust air outlet must be screened from the inside.

7. If steam is used for boosting temperature, pipes can be run the length of the floor along the
side walls discharging outwards. Steam can also be discharged directly into the air duct after
the fan. High output electric space heaters can also be used.

Filling Procedures

Depending on the growing system chosen, the compost is loaded into trays, shelves or a bulk

Compost Preparation/99

Jtr^«j^--*K>-ss hm*m • ... 1 __

Figure 96 Small Phase II room designed for trays or bulk fill

room. Certain basic principles should be adhered to when filling. These are:

1 . Fill the room as quickly as possible to minimize heat loss from the compost.

2. Compress a long strawy compost and fill loosely a short dense compost.

3. If the compost appears dry, water lightly and evenly during filling. If water streams out when a
handful is squeezed, don’t fill. Add again as much gypsum, turn and wait a few days.

4. Fill all shelves and trays evenly and to the same depth. Avoid creating pockets of compact
compost. Keep all compost within the container. No compost should hang over the sides.

5. Once finished, the floor should be cleaned of all loose compost, then washed with water.

Depth of Fill

Up to a point there exists a direct relationship between the amount of compost filled per square
foot and yield. In a fixed shelf system, the amount of compost filled is usually the amount available
for cropping. This normally holds true for trays, although some systems empty the trays at spawning
and then refill 25% fewer trays than the number that was originally filled. This results in high dry
weight efficiencies without the complications of deep compost layers during Phase II. As a general
rule, a fill depth of 8 inches will provide sufficient nutrients as well as contribute to the ease of Phase
I. At depths over 8 inches temperature stratification will lead to varying conditions within the com-

100/The Mushroom Cultivator

post, complicating the Phase II program. At depths under 5 inches there is insufficient mass for
proper heat generation and large quantities of steam may be needed.

An important consideration is the ratio of cubic feet of compost filled to cubic feet of air space in
the room. This ratio largely determines whether a supplementary heating source is necessary.
Clearly, greater volumes of compost require less additional heating. To maximize compost heat
generation, some tray systems stack trays no more than 3-4 inches apart during Phase II. These
trays are later distributed to two cropping rooms with a spacer inserted between the trays to facilitate
picking.

PAY PHASE il PROCEDURE: TRAYS OR SHELVES

0 The house is filled and cleaned. Thermometers are placed in the center of at least four
containers, and one in the middle of the room for reading air temperature. Shut the

Figure 97 Phase il temperature profile for trays or shelves.

Compost Preparation/ 101

door, turn on the fan and close the fresh air vent. Air and compost temperatures should
rise from microbial activity. If not, additional heat should be supplied. Once the compost
reaches 1 20 °F., the fresh air vent should be opened and regulated to maintain compost
temperatures in the 125-1 30 °F. range. From this point on, the fresh air vent should
never be less than the minimum setting of 10%.

1-2 A temperature chart should be kept, noting air and compost as well as fresh air and
steam settings. Temperatures should be read every 4-6 hours. Compost temperatures
should be in the 1 25-1 30 °F. range for the first 48 hours after fill. After this period, pas-
teurization should commence. The air temperature is boosted to 140°F. and held long
enough to subject the compost to 140°F. for 2 hours. If 140° can not be reached, a
compost and air temperature of 135° for four hours is sufficient. The temperatures
should be monitored closely to be sure pasteurization is complete. A long stemmed ther-
mometer can be pushed through a drilled opening in the door, or a remote reading ther-
mocouple can be used. After pasteurization, full fresh air is introduced to stop rising
compost temperatures. Once the compost temperature begins to drop, adjust the fresh
air setting to stop the compost in the temperature zone required, 128-1 30 °F.

2-10 Starting at 128°F., use fresh air to lower the compost temperature gradually, 2° per
day, until 1 22 0 is reached. Hold the compost at that temperature until it is free of ammo-
nia. Throughout this conditioning process, a compost to air differential of 10-30°F. is
normal. This differential is important for the passage of air through the compost. Little or
no differential is undesirable and indicates over-composting or under-supplementation.
During the conditioning period definite changes in the compost become apparent. The
compost becomes well flecked with whitish actinomycetes, and on the surface whitish
grey aerial mycelia of Humicola species appear. Both are indicators of proper microbial
conversion.

5-1 0 Once the compost is free of ammonia, full fresh air is introduced, dropping the compost
temperature rapidly to spawning temperatures in the 76-80 °F. range.

Phase SS in Bulk

For many people, equipping a standard Phase II room for trays or shelves may be inappropri-
ate, especially if steam is used. The recent development of the bulk system now gives the home
grower the ability to perform the Phase II without steam. This system utilizes compost heat more ef-
ficiently by loading the compost in mass, five feet deep, into a small well insulated room with a
slatted floor. Instead of air diffusing through the compost by convection, air is blown under the floor
and forced up through the compost. The wide compost to air temperature differential so essential to
conventional Phase II processes is eliminated; compost and air temperatures are now no more than
5 °F. apart. This narrow differential is in part related to a reduced compost-to-air volume ratio, which
in a bulk room is 1 : 1 or 1 This reduction of air space, coupled with the airtight, well insulated
room, results in full utilization of compost heat generation. A large measure of control over compost

102/The Mushroom Cultivator

temperatures becomes possible and optimum temperatures within the mass can be tightly regu-
lated.

Bulk Room Design Features

The size of the bulk room varies according to individual needs, but should be large enough that
there is sufficient compost mass to supply heat.

1 . At a fill depth of 4-5 feet, one ton of compost requires approximately 8-10 sq. ft. of floor

space.

2. Bulk rooms are well insulated. The walls and door are R-19; the ceiling is R-30 mini-
mum-. A vapor barrier should protect all insulation.

3. The room has a double floor. The bottom floor is concrete, insulated to R-19 with styro-
foam or other water impervious material, and covered with tar or temperature resistant
plastic as a vapor barrier. The compost floor is 12-18 inches above the bottom floor, and
is made of 4 x 4’s with spacers in between to leave 20% air space. This floor is removable
to permit periodic cleaning.

4. The interior walls and ceiling are made of exterior grade plywood, treated with a wood
preservative or marine epoxy. Allow !4 inch for expansion. Caulk or seal with fiberglass
tape.

5. The room must be airtight. Caulk all cracks and corners.

6. The access door runs the width of the room for easy loading and unloading. An airtight
seal is essential.

7. A wood plank wall is inserted before the access door to prevent the compost from press-
ing against it. The plank wall is held in place by runners on either wall.

8. The ventilation system is powered by a centrifugal, high pressure belt driven blower, with
a capacity of 90-1 20 CFM per ton of compost at a static pressure of up to 4 inches of wa-
ter gauge. The recirculation duct comes out on the top of the back wall and down to the
fan. The supply duct goes from the fan to the air chamber under the compost floor. All
ductwork should be insulated.

9. The fresh air inlet and damper are located before the fan. This damper also regulates the
recirculated air. The fresh air should be filtered.

1 0. The exhaust outlet is located on the access door. This is a free swinging damper that oper-
ates on room pressure. This outlet is covered by a coarse filter.

1 1 . Standard inside dimensions are 6-12 feet wide by 8-10 feet high.

1 2. For better temperature control the bulk room should be built inside a larger building, like a
garage, where temperaure differences are less extreme. The introduction of cold fresh air
hampers the process by neutralizing the compost heat.

A simple variation of this bulk room is a well insulated bin. The bin is constructed using the

Compost Preparation/ 103

principles just outlined. Rather than a mechanical air system, fresh air is admitted through adjustable
vents at floor level and exits through similar vents in the ceiling. Because air passage is by convec-
tion, the compost should be filled loosely and to a depth of no greater than four feet.

Figure 98 Bulk pasteurization room. Ventilation system on end wall. (Design— Vedder)

Figure 99 Bulk pasteurization room. Ventilation system on side wall. (Design — Claron)

104/The Mushroom Cultivator

Bulk Room Filling Procedures

1 . Fill as quickly as possible to minimize heat loss.

2. Compost should have good structure and optimum moisture content. Do not fill a dense,
overwet compost.

3. Fill evenly. Compost density is important. Avoid localized compaction as well as gaps.
Gaps or holes in the compost become air channels to the detriment of the surrounding
material. Be sure the compost presses firmly and evenly against all sides of the room.

4. Before filling the last three feet, put the inside board wall in place. Now fill the remaining
area. The compost should press firmly against the board wall.

DAY BULK ROOM PHASE II PROCEDURES

0 Filling. Compost is brought into the room. If remote reading temperature sensors are

used, place 2-4 sensors in different locations within the compost, and one in the air
above. If remote sensors are not used, place one thermometer in the return air duct and
one downstream from the fan in the supply duct. The compost temperature should be
within the readings of these two air thermometers. Turn the fan on, close the fresh air
damper and re-circulate until 120°F. is reached. This should take 8-24 hours. Then
open the fresh air damper to the minimum setting, 8-10%.

1- 2 Pasteurization: Allow the temperature to rise to 132-135°F. Adjust the fresh air

damper to hold this temperature for at least six hours and a maximum of ten hours.
Once completed, introduce sufficient fresh air to bring the temperature down to 122°F.
This should take approximately 1 2 hours. Be sure to anticipate temperature trends and
adjust the fresh air accordingly.

2- 10 Conditioning: By adjusting the amount of fresh air, the compost is held in the

1 18-1 22 °F. range until all ammonia is gone. Fresh air should gradually be reduced as
thermogenesis subsides. The temperature in the return air duct should always be higher
than in the supply duct.

4-10 Cool-down: Once the ammonia content of the air is below 10 parts per million (ppm)
full fresh air is given to reduce the compost temperature to 80 °F. The cool-down should
proceed as rapidly as possible.

Testing for Ammonia

The basic ammonia detection test has always been the sense of smell. The odor of ammonia
must be completely gone from the compost before it can be spawned. Odors are always good indi
cators of compost suitability. Flowever, to be absolutely certain, other methods are also used.

1 . Cresyl Orange and filter paper: Pre-cut strips of white filter paper are saturated with a few
drops of cresyl orange liquid which turns the white paper yellow. Expose the paper to the
inside of the Phase II room or to the exhaust air of the bulk room. The paper can also be

Compost Preparation/ 105

placed into small holes dug into the compost. The presence of ammonia turns the paper
varying shades of red. Purple indicates the highest concentration, while pink indicates a
lower one. When the yellow paper remains unchanged in color, free ammonia is absent.

2. Air samplers using gas detection tubes: These tubes are filled with chemicals that change
color as air samples are drawn into them. The tubes are calibrated in parts per million (ppm)
and give accurate readings down to 1 ppm. The air samplers are manufactured by Mine
Safety Co. and the Draeger Corp. Individual tubes cost from $2.00-$4.00 in lots of ten.
(See sources in Appendix).

Aspect of the Finished Compost

The following guidelines can be used to determine whether a compost is ready for spawning.
(See Color Photographs 5-8).

Figure 100 Bulk room Phase II temperature profile. (Dutch procedure)

106/The Mushroom Cultivator

1 The raw pungent odor is gone; the odor is now light artd pleasant, even slightly sweet.

2. The ammonia odor is completely gone. The cresyl orange test shows no reaction. Detector
tubes read 1 O ppm or less.

3. The pH is below 7.8, preferably 7.5.

4. Straws appear dull and uniformly chocolate brown, speckled with whitish actinomycetes.

5. The compost is soft and pliable and can be sheared easily.

6. When squeezed the compost holds its form. No water appears and the hand remains
relatively clean.

7. Moisture content is 64-66% for horse manure and 67-68% for synthetics.

8. Nitrogen content is 2. 0-2. 3%; the C:N ratio is 17:1

ALTERNATIVE COMPOSTS AND
COMPOSTING PROCEDURES

Sugar Cane Bagasse Compost

Sugar cane bagasse is the cellulosic by-product of sugar cane after most of the sugars have
been removed. It is generally a short fibrous material with a high moisture holding capacity. Total ni-
trogen amounts to 0.1 8%. In 1 960, Dr. Kneebone of Pennsylvania State University reported grow-
ing Psilocybe aztecorum on a bagasse based compost. He later reported in more detail on experi-
ments using bagasse compost for growing Agaricus brunnescens. Bagasse used as stable bedding
produced yields comparable to the horse manure based control. Bagasse supplemented with a
commercial activator (“Acto 88”) yielded poorly.

Dr. Kneebone’s composts were prepared using the standard techniques elucidated in this chap-
ter with a turn schedule on days 0-2-5-7-9. The supplemented bagasse was composted 3 days
longer and all bagasse based composts had moisture contents ranging from 75-83%. Significantly,
the bagasse compost with the lowest moisture content had the highest yield. All bagasse composts
had larger mushrooms than the control.

This work by Kneebone demonstrates the value of bagasse as a mushroom growing substrate.
Using the compost formula format, composts can be devised to meet the needs of the two species
named and many others. A good supplement would be horse droppings on wood shavings. If the
bagasse compost becomes too short or wet, the gypsum can be increased from 5% to 8% of the
dry weight.

The 5-Day Express Composting Method

During the past 20 years compost research has been directed towards shortening the overall
preparation period. The goal is to reduce handling and further conserve the nutrient base (dry mat-

108/The Mushroom Cultivator

1 1 0/The Mushroom Cultivator

T he use of non-composted and semi-composted materials as mushroom growing substrates is
common among commercial growers of Pleurotus, Volvariella, Flammulina and Stropharia.
Because of the simplicity and ease by which they are produced, these substrates are ideal for the
home cultivator. The advantages of these substrates are the rapid preparation times and the easily
standardized mixtures formulated from readily available raw materials. These substrates can be
treated by sterilization, pasteurization or used untreated in their natural state.

NATURAL CULTURE

For most people mushroom cultivation implies an indoor process employing sterile culture
techniques and a controlled growing environment. Although this has been the natural progression
of events for commercial cultivators and is the only way to consistently grow year round crops, it
need not be the sole method available to the home cultivator. For hundreds of years home growers
have made up outdoor beds and have enjoyed harvesting seasonal crops of mushrooms. In fact,
most mushrooms now being grown commercially were originally grown using natural culture tech-

niques.

By observing wild mushrooms fruiting in their natural habitats, one can begin to understand
their growth requirements. To fully illustrate how this methodology works, the development of natu-
ral culture for Psilocybe cyanescens will be used as an example. Psilocybe cyanescens grows along
fence lines and hedge rows, in tall rank grass, in berry thickets, in well mulched rhododendron beds,
in piles of wood chips and shavings and in ecologically disturbed areas. In many instances, the
mushrooms are found growing in soil, but upon dose examination of the underlying mycelial net-
work, it is apparent that they are feeding on wood or other similar cellulosic material. Due to the
thick'strandy mycelium of Psilocybe cyanescens, it is relatively easy to locate and gather colonized

Figure 102 Virgin spawn: Psilocybe cyanescens mycelium on a wood chip.

Non-Composted Subtrates/1 1 1

pieces of substrate. These pieces are considered virgin spawn and are used to inoculate similar
materials. Freshly cut chips of alder, maple and fir all support healthy mycelial growth. Because
alder is high in sugar content, without resins and abundant in northwestern North America, it has
been selected as the primary substrate material.

Even though such a virgin spawn is not absolutely clean, Psitocybe cyanescens mycelium col-
onizes fresh substrate pieces so rapidly that there is little risk of contamination. In order to prepare
inoculum for the following year, the newly inoculated chips are kept indoors in gallon jars or other
protective containers. With sufficient moisture, minimal air exchange and normal indoor tempera-
tures, the mycelium soon spreads throughout the fresh chips. For the best results a 1 :5 ratio of
virgin spawn to fresh chips is recommended. As one jar becomes fully permeated, it can be used to
produce more spawn.

In the spring freshly cut wood branches are chipped, then mixed with the fully colonized inocu-
lum and made into a ridge bed directly on the ground. Experience has shown that irregular chips
approximately 1 -3 inches long give better results than finely ground material such as sawdust. Fresh
chips not only provide a greater nutrient and water reservoir, but also have substantial surface area
for primordia formation. Strong mycelial growth can be sustained on wood chips for a prolonged
period of time. (Mycelial growth on fresh sawdust is at first rapid and rhizomorphic but soon slows
and loses its vitality).

The ridge beds should be made 4-6 inches deep and 2 feet wide. To insure a humid microcli-
mate for mushroom development the bed should be made under rhododendrons or other leafy or-
namentals, along a fence or hedge row, or on grass which is allowed to grow up through the bed.

Figure 103 Chipping freshly cut alder
branches.

112/The Mushroom Cultivator

The bed must never be placed where it is exposed to direct sunlight but it should not be so well pro-
tected that rainfall can not reach it.

During the spring and summer the mycelium colonizes the fresh substrate which should be
covered with plastic or cardboard to prevent drying. A weekly watering helps to keep the moisture
content high. In the fall the bed is uncovered and given a heavy watering twice a week, but with care
not to flood it. When the mushrooms begin to fruit, watering should be gauged according to envi-
ronmental conditions and natural precipitation. As long as the temperature stays above freezing the
mushrooms will grow continuously. If a freeze is expected, the beds can be protected with a plastic
covering. Extended freezing weather ends outdoor cropping until the following year.

Throughout the winter the beds can be protected by a layer of straw, cardboard or new chips
topped with plastic. This is particularly important for harsh climates. Other possibilities include mak-
ing the bed inside a cold frame or plastic greenhouse. Certain regions of the country like the North-
west are better suited to natural culture than others. In this respect it is desirable to use a local strain
adapted to local conditions. In climates unsuited to outdoor cultivation, the wood chips can be filled
into trays and brought inside.

Once the primary bed has been established outdoors, it can be likened to a perennial plant,
which is the nature of mushroom mycelium. Indoor spawn preparation and incubation become un-
necessary. With each successive year chips can be drawn from the original bed and used as inocu-
lum. This means that the total bed area can be multiplied by five on an annual basis. (See Figure
164 of Psilocybe cyanescens fruiting indoors in tray of alder chips).

Figure 104 Oak dowels before and after colonization by
shiitake (Lentinus edodes) mycelium.

Non -Com posted Subtrates/113

Figure 105 Shiitake plug
inserted into oak log.

Figure 106 Stacked ar-
rangement of shiitake logs in
a greenhouse.

Figure 107 Shiitake cul-
ture outdoors under shade

114/The Mushroom Cultivator

SEMI-STERILE AND STERILE
WOOD BASED SUBSTRATES

Mushrooms that grow on wood or wood wastes are termed lignicolous due to their abililty to
utilize lignin, a microbial resistant substance that constitutes the heart wood of trees. The main com-
ponents of wood, however, are cellulose and hemicellulose, which are also nutrients available to lig-
nin degrading mushroom mycelium. The chart appearing below shows a typical analysis of different
wood and straw types. This table not only illustrates the similarities between wood and straw, but
also the important differences between coniferous and broad leaf trees. The high concentrations of
resins, turpentine and tanins make conifers less suitable for mushroom growing. Conifers are used
on occasion, but they are mixed one to one with hardwood sawdust. In general, the wood of broad
leaf or hardwood species have proven to be the best mushroom growing substrates. Specifically
these tree types are: oak; elm; chestnut; beech; maple; and alder.

Hemi-

Type

Resin

P-2 0-5

K20

cell.

Cell.

Lignin

Spruce

(Picea excelsa)

2.30

0.08

0.02

0.10

11.30

57.84

28.29

Pine

(Pinus silvestris)

3.45

0.06

0.02

0.09

1 1 .02

54.25

26.25

Beech

(Fagus silvatica)

1.78

0.13

0.02

0.21

24.86

53.46

22.46

Birch

(Betula verrucosa)

1.80

—

— ■'

—

27.07

45.30

19.56

Wheat Straw

0.00

0.60

0.30

1.10

—

36.15

16.15

(Triticum sativum)

Table of the analyses of various types of wood and straw. Figures are percent of dry weight.
(Adapted from H. Rempe (1953)).

The most notable commercial species grown on wood is Lentinus edodes, the shiitake mush-
room. Traditional methods use oak logs, 3-6 inches in diameter and three feet long, cut between fall
and spring when the sap content is the highest. Special care should be taken not to injure the bark
layer when cutting and handling the logs. The bark is of critical importance for fruiting and is one of
the key factors considered by commercial growers when selecting tree species. The logs should be
scraped clean of lichens and fungi and then drilled with four longitudinal rows of one inch deep
holes spaced eight inches apart. Next, these holes are plugged with spawn and covered with wax.
After 9 to 15 months of incubation the logs begin to fruit. (See the species parameter section in
Chapter XI.) The use of freshly cut logs provides a semi-sterile substrate with no special treatment
and is a very effective method for the home cultivator.

Commercial growers of lignicolous mushrooms are turning increasingly to sawdust based
substrates. Such substrates have been developed in Japan for growing Pleurotus, Flammulina and

Non-Composted Subtrates/1 15

Auricutaria. They are also being utilized with some modifications by commercial shiitake growers in
the United States. The development of these mushroom specific substrates follows certain well de-
fined guidelines.

The basic raw material is cellulose, a major constituent of sawdust, straw, cardboard or paper
wastes, wood chips, or other natural plant fibers. Any of these materials should be chopped or
shredded, but never so finely as to eliminate their inherent structural qualities. This cellulosic base
comprises approximately 80% of the total substrate mixture.

To these basic substrate materials are added various nutrient supplements and growth stimula-
tors in meal or flour form. By supplying proteins, carbohydrates, vitamins and minerals, the supple-
ments serve to enhance the yield capabilities of the substrate base. Protein sources include concen-
trates like soya meal or soya flour, wheat germ and brewer’s yeast. The most suitable carbohydrate
sources are starchy materials such as rice, potatoes, corn and wheat. Some supplements are well
balanced and provide both carbohydrates and proteins. Examples of these are bran, oatmeal and
grains of all types. The number of possible supplements is extensive and need not be limited to
those listed. The supplements comprise approximately 8-25% of the total dry weight. The addition
of gypsum at a rate of 5% of the dry weight can improve the structure and porosity. It should be
considered an optional ingredient.

Japanese growers of Flammulina velutipes, Auricularia auricula and allies, and Pleurotus
ostreatus have a standard substrate formula consisting of 4 parts sawdust and 1 part bran. The saw-

Figure 108 Photograph of shiitake mushrooms growing on a sawdust block.

1 16/The Mushroom Cultivator

dust can be aged up to one year, which is said to improve its moisture holding capacity. Presoaking
the sawdust prior to mixing in the bran is an effective way to achieve the required 60% moisture op-
timum. A firm squeeze of the mixture should produce only a few drops of water between the fingers.
If the mixture has too much moisture, loose water collects in the bottom of the substrate container, a
condition predisposing the culture to contamination.

The substrate can be filled into a number of different containers. Mason jars, polypropylene jars
or high density, heat-resistant polyethylene bags are commonly employed. The containers are
closed and sealed with a microporous filter. They are sterilized at 15 psi for 60-90 minutes. After
sterilization the containers are cooled to ambient temperature and inoculated. The inoculum can be
either grain spawn or sawdust-bran spawn.

During incubation substrate filled plastic bags can be molded to the desired cropping form.
Common shapes are round mini-logs or rectangular blocks. Some Pleurotus growers mold the

Figure 109 Flammulina velutipes, the
Enoke Mushroom, fruiting in mason jar
containing sawdust mixture.

Figure 1 1 0 Autoclavable plastic bag and
microporous filter disc, known in the
Orient as the Space Bag.

Non-Composted Subfrates/1 17

sawdust substrate into a cylindrical shape, 6-8 inches long and 4-5 inches in diameter. The fully col
onized “logs” are stacked together on their sides with the ends exposed as the cropping surfaces.
An alternative is to slit the bag lengthwise in four places, exposing the substrate to air while retaining
the plastic as a humidity hood. If growing in jars, Flammulina and Pleurotus fruit from the exposed
surface at the mouth of the jars.

In commercial mushroom production one of the most frequently used substrate materials is
cereal straw. Not only does straw form the basis for mushroom composts, but it is also used uncom-
posted as the sole ingredient for the growth of various mushroom species. Although all types of
straw are more or less suitable, most growers use wheat because of its coarse fiber and its availabili-
ty. The straw should be clean, free from molds and unspoiled by any preliminary decomposition.
Preparation simply involves chopping or shredding the dry straw into 1 -3 inch pieces. This can be
done with a wood chipper, a garden compost shredder or a power mower. The shredding increases
moisture absorption by expanding the available surface area. Shredding also increases the density of
the substrate mass.

Figure 111 Equipment needed for pasteurization of straw: 55 gallon drum; gas burn-
er; shredder; hardware cloth basket and straw.

118/The Mushroom Cultivator

Figure 112 Shredding the straw.

Figure 113 Filling the shredded straw into the
wire basket.

Figure 114 Checking the water temperature.
Figure 115 Draining the pasteurized straw.

3TV

/ /

jLa. “

ili:.

Non-Composted Subtrates/ 119

The chopped straw is treated by pasteurization which can be carried out with live steam or hot
water. Presoaked to approximately 757o water, the straw is filled into a tunnel or steam room as de-
scribed in the composting chapter. It is steamed for 2-4 hours at 1 40-1 50 °F., then cooled to 80 °F.
and spawned. An alternative program calls for 12-24 hours at 122°F. after the high temperature
pasteurization. This program is designed to promote beneficial microbial growth giving the straw a
higher degree of selectivity for mushroom mycelium.

The method best suited to the home cultivator is the hot water bath. Figure 1 1 1 illustrates a
simple system utilizing a 55 gallon drum and a propane burner. The drum is half filled with water
that is then heated to 1 60-1 70°F. Chopped dry straw is placed into the wire mesh basket and sub-
merged in the hot water. (A weight is needed to keep the straw underwater.) After 30-45 minutes
the straw is removed from the water and allowed to drain. It is very important to let all loose water
run off.

Once drained, the straw is spread out on a clean surface and allowed to cool to 80 ° F. (or less),
at which point it can be spawned. The straw is evenly mixed with spawn and filled into trays, shelves
or plastic bags. Some compression of the straw into the container is desirable because the cropping
efficiency will be increased.

The use of plastic bags is a simple and efficient way to handle straw substrates. A five gallon bag
(1 -2 mils thick) is well suited to most situations. Two dozen nail sized holes equally spaced around
the bags provide aeration. Upon full colonization, the mycelia of species like Pleurotus ostreatus

Figure 117 Pasteurized straw stuffed into plastic bags which are then perforated with

nail size holes.

and Psilocybe cubensis actually hold the straw together, at which time the bag can be completely
removed. Another alternative is to perforate or strip the bag from the top or side to allow easy crop-
ping.

Wheat straw prepared and pasteurized in this manner can be used to grow Pleurotus ostreatus,
Stropharia rugoso-annulata, Panaeolus cyanescens and Psilocybe cubensis. It is quite possible that
other species can utilize this substrate or a modification of it. Studies with Pleurotus ostreatus have
demonstrated yield increases with the addition of 20% grass meal prior to substrate pasteurization.
Supplementation of the straw after a full spawn run is another method of boosting yields (See
Chapter VII.) Bono (1978) obtained a yield increase of 85% with Pleurotus flabellatus by adding
cottonseed meal to the fully colonized straw. The optimum rate of addition was 1 32 grams per kilo-
gram of dry straw (approximately 22 grams crude protein per kilogram straw). Bono also found that
supplementation increased the protein content and intensified the flavor of the mushrooms.

Spawning and Spawn Running in Bulk Subtrates/121

1 IvlB l]ll

M ft!

l! ■

IHillJ

i H ^ ’

Jp-yS

122/The Mushroom Cultivator

nzrr

T he inoculation of compost or bulk substrates is called spawning. The colonization of these
substrates by the mushroom mycelium is known as spawn running. At spawning and during
spawn running there are several factors that must be considered if yields are to be maximized. These
factors are:

1 . Moisture content of the substrate.

2. Temperature of the substrate.

3. Dry weight of the substrate per square foot of cropping surface.

4. Duration of spawn running.

Moisture Content

Mushroom mycelium does not grow in a substrate that is either too dry or too wet. A dry sub-
strate produces a fine wispy mycelial growth and poor mushroom formation because the water es-
sential for the transport and assimilation of nutrients is lacking. On the other hand, an over-wet sub-
strate inhibits mycelial growth and produces overly stringy mycelia. Controlled experiments with
Agaricus brunnescens grown on horse manure composts have shown yield depressions when the
moisture content deviates more than 2% from the optimum. Deviations greater than 5% generally
result in a spawn run that does not support fruitbody production. A dry compost at spawning should
be lightly watered and mixed well to guard against the formation of wet spots. For an over-wet com-
post the common procedure is to add gypsum until the loose water is bound.

Substrate Temperature

Since mushroom mycelium grows within the substrate, the substrate temperature must be
monitored closely. Thermometers are placed both in the center of the substrate— the hottest region
—and in the room’s atmosphere. These two thermometers establish a temperature differential. If the
hottest point in the substrate is 80° F. and the air is 70° F. then the temperature of the total mass
must lie within this range.

The optimum temperature for mycelial growth varies depending on the mushroom species.
Agaricus brunnescens grows fastest at 77 °F. whereas Psilocybe cubensis prefers 86 °F. Tempera-
tures higher or lower simply slow mycelial growth. The growth curve shown in Figure 1 19 illus-
trates the effect of temperature on the growth of Agaricus brunnescens mycelium. Note that growth
slows at a faster rate as the temperature rises above the optimum. Therefore the object during spawn
running is to keep the substrate within the temperature range that is optimal for the fastest growth of
mycelium.

Dry Weight of Substrate

Other factors aside, the dry weight of substrate per square foot of cropping surface largely deter-
mines total yield. Commercial Agaricus growers aim for at least five pounds of dry weight of com-
post per square foot and sometimes compress up to eight pounds per sq. ft. into their containers.
Cropping efficiencies are calculated by dividing total yield per square foot into the dry weight of one

Spawning and Spawn Running in Bulk Subtrates/ 1 23

i — P— — — jW BWi — — 1W^ ^

square foot of the substrate. Thus a yield of four pounds per sq. ft. of freshly picked mushrooms di-
vided by five pounds dry weight of substrate equals an 80% cropping efficiency. Efficiencies of
80-100% are considered to be close to the maximum yield potential of Agaricus brunnescens.

The actual amount of substrate that can be compacted into one square foot of growing area and
managed depends upon the cooling capabilities of the control system as well as the outside temper-
ature. Experiments using tracer elements in mushroom beds three feet deep have shown that nutri-
ents from the farthest point are transported to the growing mushrooms. Yields per sq. ft. increased
although at a lower substrate efficiency.

During spawn running the metabolism of the growing mycelium generates tremendous quanti-
ties of heat. Substrate temperatures normally reach a peak on the 7th-9th days after spawning and
can easily reach 90 °F. At this temperature thermophilic microorganisms become active, thereby in
creasing the possibilities of further heat generation. The substrate can easily soar above 1 00 °F. and
a compost can actually rise again to conditioning temperatures. Temperatures between 95-1 10°F.
can kill the mycelium of many mushrooms. Even if the mycelium is not completely killed, these
temperatures do irreversible harm to mycelial vitality and fruiting potential. These elevated tempera-
tures also stimulate the activity of competitor molds and may render the substrate unsuitable for fur-
ther mushroom growth. Because of the enhanced heat generating capabilities of deeply filled beds,
Agaricus growers rarely fill more than 12 inches of compost into the beds.

Figure 119 Growth curve of Agaricus brunnescens on compost.

1 24/The Mushroom Cultivator

The decision on how deep to fill the spawned substrate is an important one. Here again, the
ratio of substrate to free air space in the growing room is significant. (See Chapter IV). An efficient
method of spawn running is to the fill trays 6-8 inches deep with compost and stack them closely to-
gether in the room. In this manner the heat generated within each tray remains controllable, while at
the same time the total compost heat will be sufficient to heat the room. Outside air temperature as
well as the capacity of the heating and cooling equipment should determine how many substrate
filled containers can be placed within a given space. Fresh air is generally used to provide cooling
except when it is warmer than the room temperature.

Duration of Spawn Run

Once colonization is complete, the substrate should be cased, or if casing is not used, it should
be switched to a fruiting mode. If spawn running is continued beyond this point, valuable nutrients
that could be utilized for production of fruitbodies will be consumed by further vegetative growth. If
for some reason the cropping cycle must be delayed, the substrate should be cooled until a more
opportune time.

Spawning Methods

Spawning methods, like spawn itself, have evolved over the years. As late as 1950 Agaricus
brunnescens growers customarily planted walnut sized pieces of manure spawn or kernels of grain
spawn in holes poked into the compost at regular intervals. Using this method spawn running was
slow, and areas far from the inoculum were more susceptible to invasion by competitors. The full
potential of grain spawn was not realized until the development of “mixed spawning”. The principle
of mixed spawning is the complete and thorough mixing of the grain kernels throughout the sub-
strate. In this manner all parts of the substrate are equally inoculated, resulting in the most rapid and
complete colonization possible.

The standard spawning rate used by Agaricus growers is seven liters/ton of compost or one
quart/8 sq. ft. If spawn is readily available and cheap, it is advantageous to use high spawning rates
which lead to more rapid colonization. It is also advantageous to break up the grain spawn into indi-
vidual kernels the day before spawning. If the spawn is fresh, the grain should break apart easily. If
the spawn can not be used when fresh, it should be refrigerated at 38 °F.

The basic principle of spawn running is the same regardless of the type of mushroom or sub-
strate. COLONIZATION MUST PROCEED AS RAPIDLY AS POSSIBLE TO PREVENT
OTHER ORGANISMS FROM BECOMING ESTABLISHED. Once the mushroom mycelium be-
comes dominant, natural antibiotics secreted into the substrate inhibit competitors. To prevent inva-
sion by competitors it is important that spawning take place under carefully controlled hygienic con-
ditions. Fungus gnats in particular must be excluded, and for this purpose a tight, well sealed work-
ing area is best. This area and all tools should be disinfected one day prior to spawning with a 1 0%
bleach solution. When using disinfectants be sure your skin is protected and avoid breathing any
fumes.

Spawning and Spawn Running in Bulk Subtrates/125

Figure 120 Psilocybe semilanceata mycelium running through pasteurized wheat
straw.

If the substrate has been filled into shelves, the spawn is broadcast over the surface and mixed in
with a pitchfork or by hand. With trays, a similar method can be used, or alternatively, the substrate
can be dumped out on a clean surface, mixed with spawn and then replaced in the trays. Substrates
from a bulk room are removed, mixed with spawn and then placed into the chosen container.

It is common procedure to level and compress the substrate to avoid dehydration caused by ex-
cessive air penetration. The degree of compression depends upon substrate structure. Long, airy
materials can be compacted more than short, dense ones. Commercial tray growers compact the
compost into the trays with a hydraulic press so that the compost surface resembles a table top. This
enables the application of an even casing layer.

Environmental Conditions

The required environmental conditions for spawn running are very specific and must be closely
monitored. Substrate temperatures are controlled by careful manipulation of the surrounding air
temperature. Heating and cooling equipment are helpful but not absolutely essential unless the out-
side climate is extreme. A well insulated room with provisions for fresh air entrance and exhaust air
exit should be adequate for most situations. The steady or periodic recirculation of room air by
means of a small fan helps to keep an even temperature throughout the room and guards against lo-
calized over-heating, especially in the uppermost containers. Humidity is extremely important at this
time and must be held at 90-100%. If the humidity falls below this level, water evaporates from the
substrate surface to the detriment of the growing mycelium. Humidification can be accomplished by
steam humidifiers or by cold water misters. If steam is used, care must be taken that the increase in
air temperature does not drive the substrate temperature above the optimal range. One common

126 /The Mushroom ^C ultivator | | m

method of counteracting drying is to cover the substrate with plastic. Be ready to remove the cover-
ing during the period of peak activity if temperatures rise too quickly.

During spawn run the mushroom mycelium generates large quantities of carbon dioxide. In
fact, it has been demonstrated that mushroom mycelium is capable of C0 2 fixation. Because of this
ability to absorb C0 2 , room concentrations of 10,000-15,000 ppm are considered beneficial and
desirable. A C0 2 level high enough to stop growth is uncommon under normal circumstances. Be-
ing heavier than air, C0 2 settles at the bottom of the room, which is yet another reason for even air
circulation within the growing environment.

Super Spawning

Super spawning is also called “active mycelium spawning” vis a vis the Hunke-Till process.
Essentially, a set amount of substrate is inoculated and colonized in the normal manner. The fully
run substrate is then used as inoculum to spawn increased amounts of a similar substrate. One
could theoretically pyramid a small quantity of inoculum into a considerable amount of fully colo-
nized substrate. This technique requires the primary substrate to be contaminant free; otherwise
contamination, not mycelium, will be propagated. The possibilities inherent in this method may be
of greater application when transferring naturally occurring mycelial colonies to non-sterile yet
mushroom specific substrates. An excellent example of this is the propagation of Psilocgbe
cganescens on wood chips. (See Chapter VI.)

Supplementation at Spawning

One of the newest advances in Agaricus culture is the development of delayed release nutrients
added to the compost at spawning. These supplements are specially formulated nutrients encapsu-
lated in a denatured protein coat. They are designed to become available to the growing mush-
rooms during the first three flushes. The application rate is 5-7% of the dry weight of the substrate.
Yield increases of Vi to 1 Ib/sq. ft. are normal. Here again, complete and thorough mixing is essen-
tial to success. Caution: these materials enrich the substrate, making it more suitable to contami-
nants if factors predisposing to their growth are present. (For suppliers of delayed release nutrients,
refer to the resource section in the Appendix).

Supplementation at Casing (S.A.C.)

SACing is another method used to boost the nutritional content of the substrate. The materials
used are soy bean meal, cottonseed meal, and/or ground rye, wheat or kafir corn grains. The fully
colonized substrate is thoroughly mixed with any one of these materials at a rate of 10% of the dry
weight of the substrate. The substrate and the supplements must both be clean and free from con-
taminants; otherwise contamination will spread and threaten the entire culture. High substrate tem-
peratures should be anticipated on the second to third day after supplementation. With this type of
nutrient enhancement yield increases of Vz-2 Ibs/sq. ft. are possible.

The Casing Layer/ 127

128/The Mushroom Cultivator

A -'overing the substrate surface with a layer of moist material having specific structural character-
wictirc is called casing. This practice was developed by Agaricus growers who found that
mushroom formation was stimulated by covering their compost with such a layer. A casing layer en-
courages fruiting and enhances yield potential in many, but not all, cultivated mushrooms.

CASING CASING CASING

cjpppipc; OPTIONAL REQUIRED NOT REQUIRED

Ag. brunnescens

Ag. bitorquis

C. comatus

FI. velutipes

Lentinus edodes

Lepista nuda

PI. ostreatus

(Florida variety)

Pan. cyanescens

Pan. subbalteatus

Ps. cubensis

Ps. cyanescens

Ps. mexicana

E-

Ps. tampanensis

S. rugoso-annulata

V. volvacea

S£

In all species where the use of a casing has been indicated as optional, yields are clearly en-
hanced with the application of one. The chart above refers to the practical cultivation of mush-
rooms in quantity. It excludes fruitings on nutrified agar media or on other substrates that produce
but a few mushrooms. Consequently, casing has become an integral part of the mushroom grow-
ing methodology.

Functions

The basic functions of the casing layer are:

1. To protect the colonized substrate from drying out.

Mushroom mycelium is extremely sensitive to dry air. Although a fully colonized sub-
strate is primarily protected from dehydration by its container (the tray, jar or plastic bag),
the cropping surface remains exposed. Should the exposed surface dry out, the myceli-
um dies and forms a hardened mat of cells. By covering the surface with a moist casing
layer, the mycelium is protected from the damaging effects of drying. Moisture loss from
the substrate is also reduced.

The Casing Layer/ 129

2. To provide a humid microclimate for primordia formation and development.

The casing is a layer of material in which the mushroom mycelium can develop an exten-
sive, healthy network. The mycelium within the casing zone becomes a platform that
supports formation of primordia and their consequent growth into mushrooms. It is the
moist humid microclimate in the casing that sustains and nurtures mycelial growth and
primordia formation.

3. To provide a water reservoir for the maturing mushrooms.

The enlargement of a pinhead into a fully mature mushroom is strongly influenced by
available water, without which a mushroom remains small and stunted. With the casing
layer functioning as a water reservoir, mushrooms can reach full size. This is particularly
important for heavy flushes when mushrooms are competing for water reserves.

4. To support the growth of fructification enhancing microorganisms.

Many ecological factors influence the formation of mushroom primordia. One of these
factors is the action of select groups of microorganisms present the casing. A casing
prepared with the correct materials and managed according te-^h0|juidelines outlined in
this chapter supports the growth of beneficial microflora.

Properties

The casing layer must maintain mycelial growth, stimulate fruiting and support continual
flushes of mushrooms. In preparing the casing, the materials must be carefully chosen according
to their chemical and physical properties. These properties are:

1. Water Retention: The casing must have the capacity to both absorb and release sub-
stantial quantities of water. Not only does the casing sustain vegetative growth, but it also
must supply sufficient moisture for successive generations of fruitbodies.

2. Structure: The structure of the casing surface must be porous and open, and remain so
despite repeated waterings. Within this porous surface are small moist cavities that protect
developing primordia and allow metabolic gases to diffuse from the substrate into the air.
If this surface microclimate becomes closed, gases build up and inhibit primordia forma-
tion. A closed surface also reduces the structural cavities in which primordia form. For
these reasons, the retention of surface structure directly affects a casing’s capability to
form primordia and sustain fruitbody production.

3. Microflora: Recent studies have demonstrated the importance of beneficial bacteria in
the casing layer. High levels of bacteria such as Pseudomonas putida result in increased
primordia formation, earlier cropping and higher yields. During the casing colonization
period these beneficial bacteria are stimulated by metabolic gases that build up in the sub-
strate and diffuse through the casing. In fact, dense casing layers and deep casing layers
generally yield more mushrooms because they slow diffusion. It is desirable therefore to
build-up C0 2 and other gases prior to primordia formation. (For a further discussion on
the influence of bacteria on primordia formation, see Appendix II.)

The selection of specific microbial groups by mycelial metabolites is an excellent ex-

130/The Mushroom Cultivator

ample of symbiosis. These same bacteria give the casing a natural resistance to competi-
tors. In this respect, a sterilized casing lacks beneficial microorganisms and has little resis-
tance to contaminants.

4. Nutritive Value: The casing is not designed to provide nutrients to developing mush-
rooms and should have low nutritional value compared to the substrate. A nutritive casing
supports a broader range of competitor molds. Wood fragments and other undecom-
posed plant matter are prime sites for mold growth and should be carefully screened out
of a well formulated casing.

5. pH: The pH of the casing must be within certain limits for strong mycelial growth. An
overly acidic or aklaline casing mixture depresses mycelial growth and supports competi-
tors. Agaricus brunnescens prefers a casing with pH values between 7. 0-7. 5. Even
though the casing has a pH of 7.5 when first applied, it gradually falls to a pH of nearly
6.0 by the end of cropping due to acids secreted by the mushroom mycelium. Buffering
the casing with limestone flour is an effective means to counter this gradual acidification.
The optimum pH range varies according to the species. (See the growing parameters for
each species in Chapter XI.)

6. Hygienic Quality: The casing must be free of pests, pathogens and extraneous debris.
Of particular importance, the casing must not harbor nematodes or insect larvae.

Materials

To better understand how a casing layer functions requires a basic understanding of soil com-
ponents and their specific structural and textural characteristics. When combined properly, the
soil components create a casing layer that is both water retentive and porous.

1. Sand: Characterized by large individual particles with large air spaces in between, sandy
soils are well aerated. Their structure is considered “open”. Sandy soils are heavy, hold
little water and release it quickly.

2. Clay: Having minute individual particles bound together in aggregations, clay soils have
few air pockets and are structurally “closed”. Water is more easily bound by clay soils.

3. Loam: Loam is a loose soil composed of varying proportions of sand and clay, and is
characterized by a high humus content.

Agaricus growers found that the best type of soil for mushroom growing was a clay/loam.
The humus and sand in a clay/loam soil open up the clay which is typically dense and closed.
The casing’s structure is improved while the property of particle aggregation is retained. The
humus/ clay combination holds moisture well and forms a crumbly, well aerated casing.

There are two basic problems with using soils for casing the increased contamination risk
from fungi and nematodes, and the loss of structure after repeated waterings. Cultivators can re
duce the risk of contamination by pasteurization, a process whereby the moistened casing soil is
thoroughly and evenly steamed for two hours at 160° F. An alternative method is to bake the
moist soil in an oven for two hours at 160° F.

The Casing Layer/ 131

38AHP

CANADIAN SPHAGHUM

ORGANIC MATTER
<&CU. FT.

Figure 122 Sphagnum peat and lime-
stone flour needed for casing.

The development of casings based on peat moss has practically eliminated the use of soil in
mushroom culture. Peat is highly decomposed plant matter and has a pH in the 3. 5-4. 5 range.
Since this acidic condition precludes many contaminants from colonizing it as a substrate, peat is
considered to be a fairly “clean” starting material. Peat based casings rarely require pasteuriza-
tion. But because peat is too acidic for most mushrooms, the addition of some form of calcium
buffering agent like limestone is essential. “Liming” also causes the aggregation of the peat parti-
cles, giving peat a structure similar to a clay/loam soil. A coarse fibrous peat is preferred because
it holds its structure better than a fine peat. In essence, the properties of sphagnum peat conform
to all the guidelines of a good casing layer.

Buffering agents are used to counter the acidic effects of peat and other casing materials. Cal-
cium carbonate (CaC0 3 ) is most commonly used and comes in different forms, some more desir-
able than others.

1 . Chalk: Used extensively in Europe, chalk is soft in texture and holds water well. Chunks
of chalk, ranging from one inch thick to dust, improve casing structure and continuously
leach into the casing, giving long lasting buffering action.

2. Limestone Flour: Limestone flour is calcitic limestone mined from rock quarries and
ground to a fine powder. It is the buffering agent most widely used by Agaricus growers
in the United States. Limestone flour is 97% CaC0 3 with less than 2% magnesium.

132/The Mushroom Cultivator

3. Limestone Grit: Produced in a fashion similar to limestone flour, limestone grit is rated
according to particle size after being screened through varying meshes. Limestone grit is
an excellent structural additive but has low buffering abilities. A number 9 grit is recom-
mended.

4. Dolomitic Limestone: This limestone is rarely used by Agaricus growers due to its high
magnesium content. Some researchers have reported depressed mycelial growth in cas-
ings high in magnesium.

5. Marl: Dredged from dry lake bottoms, marl is a soft lime similar to chalk but has the con-
sistency of clay. It is a composite of clay and calcium carbonate with good water holding
capacity.

6. Oyster Shell: Comprised of calcium carbonate, ground up oyster shell is similar to lime-
stone grit in its buffering action and its structural contribution to the casing layer. But
oyster shell should not be used as the sole buffering agent because of its low solubility in
water.

Table Comparing Casing Soil Components

Absorption Potential

M a f er j a | milliliters water/ gram % Water at Sa

Vermiculite 5.0 84%

Peat 2.5 79%

Potting Soil 0.7

Loam 0-5 25%

Chalk 0-8 37 %

Limestone Grit 0.2 15%

Sand

Values vary according to source and quality of material used. Tests run by the authors.)

Casing Formulas and Preparation

The following casing formulas are widely used in Agaricus culture. With pH adjustments they
can be used with most mushroom species that require a casing. Measurement of materials is by
volume.

FORMULA 1 FORMULA 2

Coarse peat: 4 parts Coarse peat: 2 parts

Limestone flour: 1 part Chalk or Marl: 1 part

Limestone grit: Zi part Water: Approximately 1-114 parts

Water: Approximately 2-214 parts

One half to one part coarse vermiculite can be added to improve the water retaining capacity
of these casing mixtures and can be an aid if fruiting on thinly laid substrates. When used, it must

The Casing Layer/ 133

be presoaked to saturation before being mixed with the other listed ingredients.

An important reference point for cultivators is the moisture saturation level of the casing. To
determine this level, completely saturate a sample of the casing and allow it to drain. Cover and
wait for one half hour. Now weigh out 1 00 grams of it and dry in an oven at 200 °F. for two to
three hours or until dry. Reweigh the sample and the difference in weight is the percent moisture
at saturation. This percentage can be used to compare moisture levels at any point in the crop-
ping cycle. Optimum moisture content is normally 2-4% below saturation. Typically, peat based
casings are balanced to a 70-75% moisture content.

Application

To prepare a casing, assemble and mix the components while in a dry or semi-dry state.
Even distribution of the limestone buffer is important with a thoroughly homogeneous mixture be-
ing the goal. When these materials have been sufficiently mixed, add water slowly and evenly,
bringing the moisture content up to 90% of its saturation level. There is an easy method for pre-
paring a casing of proper moisture content. Remove 10-20% of the volume of the dry mix and
then saturate the remaining 80-90%. Then add the remaining dry material. This method brings
the moisture content to the near optimum. (Some growers prefer to let the casing sit for 24 hours
and fully absorb water. Prior to its application, the casing is then thoroughly mixed again for even
moisture distribution).

At this point apply the casing to the fully run substrate. Use a pre-measured container to con-
sistently add the same volume to each cropping unit.

1 . Depth: The correct depth to apply the casing layer is directly related to the depth of the
substrate. Greater amounts of substrate increase yield potential which in turn puts more
stress on the casing layer. Prolific first and second flushes can remove a thin casing or
damage its surface structure, thereby limiting future mushroom production. A thin casing
layer also lacks the body and moisture holding capacity to support large flushes. AS A
GENERAL RULE, THE MORE MUSHROOMS EXPECTED PER SQUARE FOOT OF
SURFACE AREA, THE DEEPER THE CASING LAYER.

Agaricus growers use a minimum of one inch and a maximum of two inches of cas-
ing on their beds. Substrate depths of six to eight inches are cased 1 !4 to 1 Zi inches
deep. Substrates deeper than 8 inches are cased 1 Vi to 2 inches deep. Nevertheless, ex-
periments in Holland using casing depths of 1 inch and 2 inches demonstrated that the
deep casing layer supported higher levels of microorganisms and produced more mush-
rooms. (See Visscher, 1 975). To gain the full benefits of a casing layer, an absolute mini-
mum depth on bulk substrates is 1 inch. For fruiting on sterilized grain, the casing need
not be as deep as for fruitings on bulk substrates. Shallow layers of grain are commonly
cased % to 1 inch deep.

2. Evenness: The casing layer should be applied as evenly as possible on a level substrate
surface. An uneven casing depth is undesirable for two reasons: shallower regions can

134/The Mushroom Cultivator

Figures 123, 124 &

125 Casing a tray of
grain spawn. First the
fully colonized grain is
carefully broken up and
evenly distributed into
the tray. As an option, a
layer of partially
moistened vermiculite
can be placed along the
bottom of the tray to
absorb excess water. If
the grain appears to
have uncolonized
kernels, cover the
container with plastic
and let the spawn
recover for 24 hours
before casing.
Otherwise, casing can
proceed immediately
after the spawn has
been laid out.

easily be overwatered, thereby stifling mycelial growth; and secondly, the mycelium
breaks through the surface at different times, resulting in irregular pinhead formation.
When applying the casing to large areas, “depth rings” can be an effective means to in-
sure evenness. These rings are fabricated out of flat metal or six inch PVC pipe, cut to
any depth. They are placed on the substrate and covered with the casing, which is then
leveled using the rings as a guide. Once the casing is level and even, the rings are re-
moved.

Although the casing layer must be even, the surface of the casing should remain
rough and porous, with small “mountains and valleys”. The surface structure is a key to
optimum pinhead formation and will be discussed in more detail in the next chapter.

Casing Colonization

Environmental conditions after casing should be the same as during spawn running. Substrate
temperatures are maintained within the optimum range for mycelial growth; relative humidity is
90-100%; and fresh air is kept to a minimum. (Fresh air should only be introduced to offset over-
heating). The build-up of C0 2 in the room is beneficial to mycelial growth and is controlled by an
airtight room and tightly sealed fresh air damper. If the entrance of fresh air cannot be controlled, a

Figure 126 Depth rings used for even casing application on bulk substrates.

136/The Mushroom Cultivator

Figure 127 Mycelial growth (Agaricus brunnescens) into casing with optimum
moisture.

sheet of plastic should be placed over the casing. This plastic sheet also prevents moisture loss from
the casing.

Soon after casing, substrate temperatures surge upward due to the hampered diffusion of
metabolic gases which would normally conduct heat away. This surge is an indication of mycelial
vitality and is a positive sign if the room temperature can be controlled. This temperature rise can
be anticipated by lowering either the temperature of the substrate prior to casing or lowering the
air temperature of the room after casing.

Within three days of application, the mycelium should be growing into the casing layer. Once
mycelial growth is firmly established, the casing is gradually watered up to its optimum moisture
holding capacity. This is accomplished by a series of light waterings with a misting nozzle over a
two to four day period (depending upon the depth of the casing). Deeper casings require more
waterings. Optimum moisture capacity should be achieved at least two days before the mycelium
reaches the surface. IT IS EXTREMELY IMPORTANT THAT THE WATERINGS DO NOT
DAMAGE THE SURFACE STRUCTURE OF THE CASING. Heavy direct watering can “pan”
the casing surface, closing all the pore spaces and effectively sealing it. The growing mycelium is
then trapped within the casing layer and may not break through it at all. The ultimate example of
panning is a soil turned to mud.

To repair a casing surface damaged by watering, the top !4 inch can be reopened by a tech-
nique called “scratching”. The tool used is simply a 1 x 2 x 24 inch board with parallel rows of

The Casing Layer/ 137

Figure 128 Mycelial growth (Psilocybe cubensis)
into casing with optimum moisture.

nails (6 penny) slightly offset relative to one another. With this “scratching stick”, the casing is
lightly ruffled prior to the mycelium breaking through to the surface. After the surface has been
scratched, the casing should be given its final waterings prior to pinning.

A modified application of this technique is “deep scratching”. When the mycelium is midway
through the casing, the entire layer is thoroughly ruffled down to the bulk substrate. The agitated
and broken mycelium rapidly reestablishes itself and within three to four days it completely colo-
nizes the casing. The result is an early, even and prolific pinhead formation. Before using this
technique, the grower must be certain that the substrate and casing are free of competitor molds
and nematodes.

Casing Moisture and Mycelial Appearance

Moisture within the casing layer has a direct effect on the diameter and degree of branching
in growing mycelium. These characteristics are indicators of moisture content and can be used as
a guide to proper watering.

1. Optimum Casing Moisture: Mushroom mycelium thrives in a moist humid casing,
sending out minute branching networks. These networks expand and grow, absorbing
water, C0 2 and oxygen from the near saturated casing. This mycelial growth is character-
ized by many thick, white rhizomorphic strands that branch into mycelia of smaller dia-
meters and correspondingly smaller, finer capillaries. The overall aspect is lush and

138/The Mushroom Cultivator

dense. When a section of casing is examined, it is held firmly together by the mycelial
network but will separate with little effort. The casing itself remains soft and pliable.

2. Overly dry casing: In a dry casing, the mycelium is characterized by a lack of rhizo-
morphs and an abundance of fine capillary type mycelia. This fine growth can totally per-
meate the casing layer, which then becomes hard, compact and unreceptive to water. It is
common for puddles to form on a dry casing that has just been watered. Also, a dry cas-
ing rarely permits primordia formation because of its arid microclimate and is susceptible
to “overlay”. Mushrooms, if they occur, frequently form along the edges of the tray.

Overlay is a dense mycelial growth that covers the casing surface and shows little or
no inclination to form pinheads. Overlay directly results from a dry casing, high levels of
C0 2 and/or low humidity. (See Chapter IX on pinhead initiation).

3. Overly Wet Casing: In a saturated casing, the mycelium grows coarse and stringy, with
very little branching and few capillaries. Mycelial growth is slow and sparse which leaves
the casing largely uncolonized. Often the saturated casing leaches onto the substrate sur-
face which then becomes waterlogged, inhibiting further growth and promoting contami-
nation. Subsequent drying may eventually reactivate the mycelium, but a reduction in
yield is to be expected.

140 /The Mushroom Cultivator

T he change from the vegetative state of mycelial growth to the generative one of primordia for-
mation is called pinning, pin setting, pinhead initiation or fructification. Primordia or pinheads
are knots of mycelium that precede development info small mushrooms. All species reguire a set of
environmental conditions for pinning that are quite different from the conditions for mycelial growth.
By understanding the factors that regulate this change in the mushroom life cycle, the cultivator can
control the pinning process.

In nature primordia formation is primarily influenced by seasonal changes in environmental
conditions. In temperate climates most mushrooms fruit during the cool, wet fall whereas in tropical
and subtropical climates mushrooms fruit during the rainy season. The fruiting period ends when
the season changes and environmental conditions become too hot, too cold or too dry. The myce-
lium then lies dormant or grows slowly, reactivated only by the warming of spring and summer.
These seasons are times for the mycelium to expand its network, absorb nutrients and rebuild its
energy reserves. Once the cool wet conditions of fall return, these reserves are used to support an-
other crop of mushrooms.

Basic Pinning Strategy

Mushrooms fruit indoors in response to much the same conditions that trigger fruiting in the
wild. Several environmental factors, working in combination, provide an ideal environment in which
mushrooms flourish. Most, if not all cultivated mushrooms fruit at lower temperatures than the opti-
mum for the growth of mycelium. Usually, a drop in temperature is accompanied by rain or an in-
crease in humidity. Water is essential for the absorption of nutrients by the mycelium. And vaporous
water creates the humid microclimate that is so critical for the developing primordia. Primordia have
a low tolerance to C0 2 and need ample fresh air. And while the mycelium has no requirement for
light, many species need light to initiate pinheads and to mature into healthy mushrooms. Mush-
rooms form only when there is a coincidence of all these factors. Cultivators create an artificial envi-
ronment that prolongs these optimum conditions so that mushrooms are given the best possible en-
vironment in which to grow.

Primordia formation strategies are well defined for species now under cultivation. These proce-
dures are similar in their approach and differ only in certain environmental requirements. Given that
the substrate has sufficient nutrients, the interaction of water, humidity, temperature, fresh air, C0 2
and light all play determining roles in the fructification process. (In some cases, specific microorgan-
isms must be present before fruiting can occur). The modification of any one of these factors be-
yond the fruiting requirements can inhibit or stop the process. Hence, the cultivator must have pre-
cise control over conditions within the growing room if this critical phase is to be carried out suc-
cessfully.

Strategies for Mushroom Formation/ 141

Agaricus brunnescens culture illustrates the interplay of environmental factors in pinhead initia-
tion. It serves as a useful model for setting primordia in many species, especially those using a cas-
ing layer. In each of the following stages, the main considerations are highlighted and then dis-
cussed in detail. Although Agaricus does not require light, and since most cultivated mushrooms
do, this requirement has been listed as the last parameter.

Stage I: Preparation

Following its application, the casing is conditioned to allow even mycelial growth into it. Once
mycelial growth is well established, the casing layer microclimate and the growing room are careful-
ly managed to meet the following requirements.

1. The casing layer is at optimum moisture capacity.

2. The casing layer surface is rough and porous.

3. The relative humidity of the growing room’s air is 95%.

4. The substrate is incubated in total darkness.

During the casing colonization period, the casing layer is being conditioned for pinhead initia-
tion. Gradually, the moisture content is brought up to the optimum and a microclimate with high
relative humidity is carefully maintained. Water in the casing moves by capillary action to the surface
where it is drawn into the air by evaporation. This constant movement slowly depletes the casing of
the moisture needed to protect pinhead development. Therefore, in conjunction with an optimum
casing moisture level, the relative humidity of the room must be held at 95%. Lower humidities
must be accompanied by light but regular waterings. The higher the humidity (rFH), the less water
will be lost to evaporation.

Given optimum moisture conditions in and directly above the casing layer, the next step is to
prepare the casing surface. Whether by initial application or by ruffling at a later time, the casing sur-
face should be rough and open— with minute mountains and valleys. A rough open casing has
more surface area where pinheads can form, provides a humid environment conducive to that for-
mation and allows the diffusion of metabolic gases.

Stage II: Environmental Transition — The Prelude to Setting Primordia

Pinhead initiation techniques should begin when the mycelium reaches the valleys of the cas-
ing surface. Once the mycelium is clearly established in the valleys, the cultivator can begin the first
steps leading to the setting of pinheads. Within this one to two day period, the

1. Substrate and air temperatures are lowered to the fruiting range.

2. The humidity is maintained at the 95% level.

3. The carbon dioxide content of the room is reduced by the introduction of fresh air.

4. The room is lighted on a 12 hour on/off cycle.

1 42/The Mushroom Cultivator

Figure 130 Overlay.

Mycelium breaking through the casing surface early should be lightly sprinkled with moist cas-
ing. Uneven growth through the casing layer is usually an indication of a casing with irregular
depths. By “patching” shallow areas, an even mycelial spread is assured. Note that the more even
the distribution of the mycelium in the valleys of the casing’s surface, the more even the
pin-set and the greater the first and second flushes.

The exact time for initiation varies with the strain and according to the experience of the individ-
ual grower. Some strains continue to grow vegetatively for a period after the initial temperature
shock whereas others stop immediately. For this reason, some cultivators initiate when 20% of the
valleys show mycelial growth while others wait until 90% are run through with mycelium. Normally
within 1 2-48 hours from the time the mycelium is first visible in the valleys, the initiation sequence
is started.

The first step in the pinhead initiation process is to lower the substrate and air temperature from
the mycelial growth optimum to the fruiting range. This temperature “shock” is accomplished by
ventilation with a large volume of cool fresh air, thereby lowering the room’s temperature to a point
5-20° below the optimum for spawn running. (For Agaricus brunnescens, this would mean drop-
ping air temperature from 70 °F. to 64 °F.). Whatever the air temperature may be, the bed tempera-
ture is normally several degrees warmer. The length of time needed to affect this change is deter-
mined by the total volume of substrate and the temperature of the air being introduced. Within 48
hours, the substrate temperature should fall to fruiting temperatures, effectively slowing vegetative
growth. This change signals to the mycelium that it is time to fruit.

Strategies for Mushroom Formation/ 143

Figure 131 Cased grain culture of Agaricus brunnescens showing overlay and stroma.

Fresh air also removes high concentrations of carbon dioxide and other metabolic gases from
the room. Since Agaricus brunnescens does not pin properly at C0 2 concentrations above 2000
ppm, lowering the carbon dioxide content of the room’s air to under 2000 ppm is critical. The in-
hibitory effect of carbon dioxide on mushroom formation gives Agaricus growers a high degree of
control over the pinning process. Not until carbon dioxide is removed will pinheads form. If carbon
dioxide levels remain high, the mycelium will totally cover the casing surface, a condition called
overlay.

The mycelial mat formed by overlay makes the casing impervious to water and produces few
pinheads. Overlay also occurs if the casing surface is too dry, the humidity (rH) is too low or the air
temperature remains too high. Overlay can be counteracted by patching, but the cause must be
diagnosed and carefully corrected if the culture is to be revived. Few flushes will be as great from a
casing with overlay as from a casing properly managed.

Stage 111: Primordia Formation (Knotting)

Once substrate temperatures have been lowered and C0 2 levels have been reduced, primordia
will begin to form. Maintain:

1. A constant fresh air supply to remove metabolic gases, and C0 2 at levels less than 1000
ppm.

2. A constant temperature in the growing room that is within the fruiting range.

144/The Mushroom Cul tivator ^

3. A relative humidity of 95%.

4. A 12 hour on/off light cycle.

The combination of temperature drop, high humidity and reduction of metabolic gases by a
constant supply of fresh air now provides an environment conducive to pinhead formation. These
parameters should be held constant until the pins are set. Any abrupt changes in temperature or hu-
midity will be harmful to primordial growth. Pinhead initials form in the humid valleys of the casing
layer and are visible as small knots of mycelium. This is the earliest stage of fruiting. Within five days
these knots enlarge into small mounds or buttons that soon differentiate into mushrooms.

Due to slowed mycelial growth in the cooled substrate, carbon dioxide evolution is greatly re-
duced. Consequently, the fresh air supply can be moderated to the minimum level necessary to
maintain 1000 ppm of carbon dioxide. At this time, oversupply of fresh air can lead to high evapo-
ration rates and excessive drying. The humidity should never be allowed to fall below 90%. If dry
air becomes a problem, a light misting of the casing surface, two to five times daily, should keep the
microclimate moist. In fact, some growers knock down the mycelium with a forceful watering on
the first day of initiation. Others mist daily as a standard practice. However, once pinning has begun,
any forceful watering will kill a number of developing pins, and damage others. Given sufficient cas-
ing moisture and a high humidity, these watering practices become unnecessary.

Stage IV: Pinhead Development

After the pinheads have grown to pea size (3-5 mm.), their further development is primarily de-
pendent on air temperature and relative humidity. To insure that they mature into healthy mush-
rooms, the

1. Air temperature is held constant within the fruiting range.

2. Relative humidity is lowered to 85-92%.

3. A constant fresh air supply with C0 2 below 2000 ppm.

4. A 1 2 hour on/off light cycle.

The humidity is lowered to 85-92%, thereby increasing the evaporation rate, an essential re-
quirement for pinhead maturation. If humidity remains too high, pinhead development will be re-
tarded. The easiest way to reduce humidify is to raise the air temperature by 1 -2 °F. or to increase air
movement within the room. Under no circumstances should pockets of stagnant air be allowed to
form. Evaporation is negligible in stagnant air pockets which are also excellent breeding grounds for
mushroom pathogens.

At this time, a slightly higher level of carbon dioxide is desirable (in the 1500-2000 ppm
range) and fresh air can be cut back accordingly. Given proper C0 2 levels, and sufficient evapora-
tion, the pins continue to develop. The exact rate of growth depends on the air temperature in the
room. Work done by Lambert (1 938) has shown that a pinhead of Agaricus brunnescens with a di
ameter of 2 millimeters fully develops into a mature mushroom in twenty-two days at 50 °F., in ten
days at 60 °F. and in six days at 70 °F. Although mushrooms develop more quickly at 70 °F., over-
all yields diminish. Optimum temperature for cropping in Agaricus brunnescens is 62-64°F.

Strategies for Mushroom Formation/ 145

Figure 132, 133 &
134 Three day
pinhead development
sequence in Agaricus
brunnescens.
Change-over from
Stage III to Stage IV
occurs within this
time frame.

146/The Mushroom Cultivator

The importance of the primordia formation period can not be over-emphasized. For maximum
yields an optimum number of pinheads must be set, matured and brought to harvest. Certain rela-
tionships exist between the pinning process and yields. These are:

1. During the primordia formation period, pinheads for the first and second flush
are being generated. The second flush primordia are present as thickened mycelial knots
which develop after the first flush is harvested. Once the first flush is off the beds, the second
set of primordia begin to enlarge and within days attain button size. Because 60-75% of the
total yield is normally harvested from the first two flushes, the few days of pinhead initiation
are the most critical in the growing of mushrooms. Hence, all environmental factors must
be carefully monitored to insure the best possible pin-set.

Strategies for Mushroom Formation/ 1 47

2. The greater the number of pins set for the first flush, the higher the yield, provided
sufficient nutrients are available to support their growth. However, with more pinheads
competing for the same nutrient base, the smaller are the mushrooms arising from it. Fewer
pinheads result in larger mushrooms, but lower total yields.

3. The substrate will only support the development of a certain number of primor-
dia per flush. Under normal circumstances with an even pin-set, pinheads may “abort”
because of insufficient nutrients or late formation.

4. Pins that form early delay the growth of neighboring primordia. Good examples of
this can be found in shallow areas or along the borders of the substrate container. Remov-
ing these relatively few “volunteers” before they develop is advantageous to the remaining
primordia that constitute the first flush.

THE INFLUENCE OF LIGHT

ON PINHEAD INITIATION

Mushroom species requiring light for primordia formation are said to be photosensitive. Al-
though light is not necessary to induce fructification in all mushrooms (i.e. Agaricus brunnescens),
certain spectra have proven to be stimulatory to pinhead initiation and are critical for the normal de-
velopment of the fruitbody. Psilocybe cubensis and Pleurotus ostreatus are two such photosensitive
species.

A thorough investigation on the photosensitivity of Psilocybe cubensis can be found in a mas-
ter’s thesis by E.R. Badham (1979). His work reinforces the conclusions of other researchers work-
ing with the Basidiomycetes: more pinheads are initiated upon exposure to blue and ultra-violet light
with distinct peaks at 370, 440 and 460 nanometers. Badham showed that light stimulation at
these wavelengths for as little as half a millisecond per day caused primordia to form. In contrast,
red, infra-red and green light having wavelengths greater than 510 nanometers were ineffective.

With this knowledge, the cultivator of photosensitive species can develop initiation strategies
incorporating the influence of light. Ideally a fully colonized substrate should be incubated in total
darkness and exposed to light only after the mycelium first shows through the casing layer. If the
cultivator wants to check the culture without the chance of premature pinning, red light is recom-
mended. (The proper location and type of light is discussed in more detail in Chapter IV).

148 /The Mushroom Cultivator

Environmental Factors/ 149

Figure 136 Wild strain of Agaricus brunnescens fruiting in bag of cased compost.

1 50/The Mushroom Cultivator

F or the home cultivator the onset of cropping is a time of excitement and anticipation. It is also a
time for increased attention to the finer details of environmental control. Temperature, humidi-
ty, light and airflow in the growing room all play vital roles which together determine the nature of
further mushroom development.

Temperature

During the vegetative growth period, the substrate was held in the optimum range by careful
manipulation of the air temperature. But once the change to generative growth is initiated, the sub-
strate temperature becomes less important and air temperature becomes the controlling factor.

The time it takes button sized mushrooms to mature is influenced primarily by the air temper-
ature of the growing chamber. Each species has an optimum temperature for fruitbody develop-
ment that lies within a broader growing range. Knowing the temperature parameters as outlined in
Chapter XI, the cultivator can speed or slow development depending on which end of the cropping
range is chosen. Lower temperatures can be used to postpone or lengthen the harvesting period
and allow for maximum quality control. High temperatures serve to shorten the cropping period by
promoting rapid, intense flushes. However, the dangers of high temperatures include the risk of
heat building up in the substrate and consequent C0 2 generation, as well as the ability of insects and
contaminants to grow and reproduce at faster rates. Commericial Agaricus growers commonly
lower the air temperature by 2 °F. 48 hours prior to the peak of the first and second flushes. Further
flushes are then run hotter to speed the crop to completion. It is important that the cultivator evalu-
ate the heat generating capabilities of the crop and insure that the environmental control system is
capable of handling them.

Flushing Pattern zs

The mushroom crop grows in cycles c^l^vfl^|hes or “breaks”. Depending on the species be-
ing grown these flushes normally come in sWAln day intervals with each successive flush bear-
ing fewer mushrooms. The manner in whiclvfllesfjflushes appear is determined during the pin initi-
ation period. Even pinning sets up a uniform pattern of flushing that continues throughout the crop-
ping cycle. Uneven flushing creates difficult situations for proper watering and environmental con-
trol. To encourage even flushing, early forming pinheads are picked off as buttons unless it appears
that these pins constitute the flush itself. Poor first flushes are indicative of faulty pinning procedures
and lead to lower total yields and a longer cropping period as the cultivator tries to maximize yields
from the following flushes. But keep in mind that many times it is the progressive build-up of com-
peting contaminant organisms that eventually bring mushroom growth to a halt. Fpr this reason, the
goal is to maximize yields in the early flushes.

To further increase the flushing speed the actual harvest period in each flush should be kept
short and concise. Late developing mushrooms are removed with or on the day after peak produc-
ion. The sooner the flush is completely removed the quicker the next one will appear and the short-

Environmental Factors/ 151

Figure 137 Agaricus brunnescens af-
fected by high C0 2 concentration. Note
long stems and underdeveloped caps.
Figure 138 The effect of dry air on
Psilocybe cubensis caps, a condition
known as “scaling”.

152/The Mushroom Cultivator

Figure 139 Rosecomb on Psilocybe Figure 140 Fruitbody abnormality occa-

cubensis, an abnormality caused by con- sionaily seen in Psilocybe cubensis.

tact with chemicals, especially those that
are petroleum based.

er the overall cropping cycle. Stunted undeveloped mushrooms are also cleared from the cropping
surface between breaks with care not to disturb the casing. Small dead pinheads should be left in
place and cause little harm. (As a rule, an aborted mushroom can be removed as long as the casing
is not touched in the process.) At no time should the casing be over-handled in an attempt to clean.
Such handling can spread disease spores and damage subsequent pin formation.

Air Movement

Air movement in the growing room is designed to create an even flow across all levels of crop-
ping surface. This even airflow counteracts temperature stratification and dead air pockets by equal-
izing the environment of the room. In this manner the crop can be managed as a whole, giving the
grower greater control over the cropping cycle.

During the pin initiation period fresh air is introduced into the room to remove metabolic gases
produced by the mushroom mycelium. Although gas production is reduced once this vegetative

Environmental Factors/ 153

growth has been slowed, the maturing mushrooms create more carbon dioxide, the removal of
which requires a continuous supply of fresh air. The number of these air changes varies depending
upon the air/bed ratio and the C0 2 requirement of the mushroom species being grown. Agaricus
bitorquis needs only half the amount of fresh air required by Agaricus brunnescens. A common
rate for Agaricus brunnescens is 4-6 changes per hour. For more C0 2 tolerant species such as
Psilocgbe cubensis, 2-3 changes per hour is sufficient. (The most accurate method for determining
fresh air requirements employs the multiple gas detector. This instrument measures C0 2 content of
the air in parts per million (ppm), from 300 (natural level) up to 20,000 ppm. See Appendix for
sources.)

Because many mushrooms are sensitive to carbon dioxide, the physical development of the
mushroom can also be used as a guide. High C0 2 environments produce long stems and small un-
derdeveloped caps in Agaricus brunnescens and Pleurotus ostreatus. Pleurotus exhibits similar
symptoms in conditions of low light intensity.

In general, too much fresh air is preferable to insufficient air supply. However, fresh air dis-
places the existing room air which is then exhausted from the room. Unless this fresh air is precon-
ditioned to meet the requirements of the species, one will be constantly disrupting the growing envi-
ronment and thereby over-working the heating and humidification systems. For this reason the air
circulation system should be designed to recirculate the room air. This is accomplished by a mixing
box with an adjustable damper that proportions fresh and recirculated air. In this regard, C0 2 toler-
ant species give the grower a distinct advantage in maintaining the correct environment because
they need less fresh air for growth.

An important effect of air circulation and fresh air supply is the evaporation of moisture from the
cropping surface. Excessive humidity without adequate air movement and evaporation retards
mushroom development. Saturated stagnant air pockets are also breeding areas for contaminants

154/The Mushroom Cultivator

like the Forest Green Mold (Trichoderma) and Bacterial Blotch (Pseudomonas). As stated in the
previous chapter on pinhead initiation, once the primordia are set, the relative humidity should be
lowered to 85-92% and held constant within this range throughout cropping. Besides the creation
of a cool surface by “evaporative cooling”, evaporation aids in the transport of nutrients (in solution)
from the substrate to the growing mushrooms. If the evaporation rate is too high and the humidity
falls below 85%, excessive drying occurs, causing small stunted mushrooms and cracked scaly
caps. A dry cropping surface further reduces yields and is difficult to recondition. In this respect, it is
critical for the grower to reach a balance between air circulation, fresh air and humidification. This is
but one aspect of the “Art” of mushroom culture.

Watering

Maturing mushrooms have water requirements that must be met if maximum yields are to be
achieved. Mushrooms grown on uncased substrates draw their moisture from the substrate, where-
as those grown with a casing draw equally from both. Uncased substrates are more susceptible to
dry air and therefore require a relative humidity of 90-95% as well as periodic misting of the crop-
ping surface. If the cropping surface dries and forms a dead mycelial mat, it can be reopened to fur-
ther flushing by raking or scratching. This technique is often used by Pleurotus growers to stimulate
later flushes.

The advantages of using a casing layer are many. Protected from atmospheric drying, the sub-
strate moisture is channeled solely to the mushroom crop. And, the water reservoir provided by the
casing not only supplies the mushroom flushes but also serves to keep a high humidity in the crop-
ping surface microclimate. In order to sustain these benefits, the grower must learn to gauge casing
moisture and know when to water.

Other than light mistings, any substantial waterings before the button stage can result in dam-
aged pins. But once the mushrooms have reached button size, it is time to begin building the casing
moisture back up to the peak reached at pre-pinning. The aim is to reach capacity just prior to the
main harvest. This is accomplished by a series of daily, light to moderate waterings with a fine mist-
ing nozzle. Commercial Agaricus growers have traditionally used a rose-nozzle but many have now
switched to nozzles with finer sprays and variable volume outputs. This enables the grower to add
moisture without damaging the casing surface. In this regard, high water pressures and close nozzle
proximity to the casing should be avoided.

The goal is to keep the surface of the casing open and porous throughout the cropping cycle.
Putting on too much water at once is the most common cause of panning. By watering 2-4 times/
day rather than just once, the casing can slowly absorb the water without damage to the surface.

After the first flush is harvested the casing should be kept moist with light mistings until the next
flush reaches the button stage. The casing moisture is then built up again. Each new flush is treated
in this manner, although later flushes will have fewer mushrooms and therefore require less water.
At no time should the casing be allowed to dry out. Mushrooms pulled from a dried casing carry
large chunks of casing with them, creating gaps in the cropping surface and at times exposing the

Environmental Factors/ 155

substrate to possible colonization by contaminants. If the substrate is exposed during picking, the
holes should be filled with moist casing. To recondition a dry casing, moisture should be added
slowly over a period of a few days

One of the common contaminants in mushroom growing is Bacterial Blotch (see Chapter
XIII). Blotch results from mushroom caps that remain wet for extended periods of time. Agaricus
growers attempt to dry recently watered mushroom caps as quickly as possible by lowering the hu-
midity of the room. This is accomplished by increasing air circulation and introducing more fresh air
or by raising the air temperature 1 -2 0 F. Agaricus growers also stop watering once the mushroom
cap has reached adolescence because wet mushroom caps become prime sites for disease. Small
scale growers may be able to water around maturing mushrooms without directly hitting the caps. If
Bacterial Blotch or other diseases appear on the mushrooms or the casing soil, these areas should
not be watered. Watering contaminated regions will spread the infection further. A common strategy
for serious disease outbreaks is to lower the relative humidity and run the casing drier than normal.
Agaricus cultivators also use slightly chlorinated water (150 ppm).

Harvesting

The way an individual picks mushrooms can dramatically affect future flushes. Damage to rest-
ing pinheads and disturbance of the casing soil must be minimized during picking. Often times pin-
heads are in close proximity to developing mushrooms and enlarge directly after the mature ones
are picked. Should any pinhead be harmed, the grower will have lost a potential fruitbody. More-
over, these damaged pinheads are easily parasitized by fly larvae and other contaminants. The best
pickers are meticulous, unhurried, and above all treat the mushrooms with care. Carelessness in
picking, when multiplied by hundreds of cultures, can be costly indeed.

The most important factor in harvesting mushrooms is timing. Agaricus brunnescens should
be picked before the veil breaks and the stem elongates. Psilocybe cubensis is morphologically dis-
tinct from Agaricus species, having a longer stem, a less fleshy cap and a more delicate veil. It is
both natural and desirable to have tall stands of Psilocybe mushrooms while this is not the case with
Agaricus. Cultivators of these two species, however, share many things in common. One particular
problem is the massive release of spores from the mature mushrooms. These spores often times
cover the casing layer and can inhibit further pinhead development. High spore loads can also
cause allergic reactions amongst workers. For these reasons, one should pick the mushrooms at the
stage when the veil begins to tear or soon thereafter.

The nature of the crop determines how the mushrooms should be picked. Flushes with mush-
rooms in varying stages of development are more difficult to harvest. This is especially true if the
primordia formation period was interrupted by fluctuations in the environment. One example is a
phenomenon common to Psilocybe cubensis culture in mason jars. Mushrooms sometimes form
between the casing and the glass. These “border breaks” are due to high humidity pockets and pre-
mature light stimulation. In tray culture where mycelium is not exposed to side light and proper
moisture is easily managed, border breaks are uncommon. Mushrooms then grow uniformly from
the surface of the casing layer where they can be easily picked.

156/The Mushroom Cultivator

Harvesting techniques:

1 . Equipped with a basket and short bladed paring knife, grasp the base of the stem, and with
a twisting motion, pull the mushroom from the casing layer being careful not to disturb
neighboring pinheads.

2. Trim the stem base, removing only flesh to which the casing or substrate is attached. Mush-
rooms having thin stems are best cleaned using a knife in a downward scraping motion. All
trimmings should be placed in a sealed plastic bag and removed from the cropping area.

3. Mushrooms growing in clumps or clusters should be broken apart and harvested individual-
ly when possible. Special care must be taken with those clumps containing both mature
and immature mushrooms. Leave immature mushrooms attached to the casing layer or
substrate to insure continued growth.

Preserving Mushrooms

If not served within four days, mushrooms can be preserved by drying,' freezing or canning. Air
drying of mushrooms is the method most widely used by home cultivators and field hunters. Since
most mushrooms are 90% water, they must be dried within a few hours or fly larvae and bacteria
will consume them. Provided mushrooms are placed in a flow of warm, dry air, this large fraction of
water soon evaporates into the air. Dried mushrooms are smaller, lighter and less fragrant than fresh
ones. Once dried, they are sealed in airtight moisture proof plastic containers and refrigerated.
Mushrooms will be preserved for years in this manner. When needed, simply rehydrate them in wa-
ter before cooking. They will regain much of their original size and flavor.

Commercially available food dehydrators are well suited for drying mushrooms. Their only dis-
advantage is that the trays are often too close together, necessitating the cutting of large mushrooms
into thin slices. Or, one can build a dehydrator solely designed for this function and customized to
an individual’s particular needs. A good dryer should be able to dry the mushrooms in 24-48 hours
by passing warm air no hotter than 1 1 0 °F. Open air drying at room temperature is also feasible us-
ing dehumidifiers in combination with air circulation fans. “Flash” drying at high temperatures
should be avoided since the mushrooms lose much of their nutritive value and, as the case may be,
much of their psilocybin content.

Freezing is another method of preserving mushrooms. But unless the mushrooms are first
dried, frozen mushrooms are soggy and unappealing upon thawing. In freezing, the water constitut-
ing 90% of a mushroom’s mass becomes crystallized. Frozen mushrooms are held together more
by ice crystals rather than their own cellular structure. Since ice expands upon crystallization, cells
break under the stress. Because frozen mushrooms disintegrate into a formless mass when thawed,
they are mostly used in soups or stews.

The best of both drying and freezing is freeze drying, This is the ideal method for preserving
the flavor, nutrition, form and/or psilocybian content of mushrooms. Because of the expense, only
a few commercial mushrooms, such as shiitake ( Lentinus edodes) are freeze dried.

Environmental Factors/ 157

Freeze dryers operate on the principle of first flash freezing fresh mushrooms which are then
placed onto heated trays in a cooled, high vacuum chamber. The frozen water within the mush-
rooms begins to melt from the heat generated from the trays. But instead of becoming a liquid, the
water is immediately transformed into a vapor that is pumped out of the freeze drier. Freeze drying
preserves much of the original cell structure and hence mushrooms dried in this manner are often
life-like in appearance. Since commercial freeze driers are prohibitively expensive, few home
cultivators can afford them. Many people have discovered, however, that mushrooms placed in a
frost-free refrigerator are almost as well preserved.

Canning is another method for storing mushrooms. Mushrooms preserved by canning must be
carefully cleaned beforehand, precooked for 3 or 4 minutes in boiling water, then inserted into glass
jars with a small amount of vinegar and sterilized in a pressure cooker. (Sterilization for mushrooms
is usually 30-40 minutes at 1 0 psi. Consult a book on mushroom cookery for further information
on canning mushrooms). Canned mushrooms, especially those that have been pickled, are pre-
ferred by many epicureans to thcssj! preserved by other means.

No matter what the tech |ii|lN&§jph mushrooms are undoubtedly better tasting than preserved
mushrooms. If one chooses Wjm/55eeze or can, young mushrooms should be selected over old
ones. Label each container with th^lpecies, the name of who grew or identified the mushrooms,
the date and the place of origin. (One general rule recommended by all mycologists is: when eating
wild mushrooms for the first time, always leave one or two small specimens aside in case illness en-
sues and a mycologist or a doctor needs to be consulted.)

160/The Mushroom Cultivator

G rowing parameters for mushrooms vary with every species. Through time spent in
countless trials and from observations by both home and commercial cultivators, specific
cultural requirements have been ascertained.

Mushrooms fruit in response to unique sets of conditions involving nutrition (substrate),
temperature, pH, relative humidity, light and carbon dioxide. What follows are outlines pin-
pointing the optimal environmental ranges for each stage in the mushroom’s life cycle. By ad-
hering to these optima, a cultivator can maximize fruitbody production in a precise and delib-
erate fashion.

Growing Parameters for Various Mushroom Species/ 161

SPECIES: Agaricus bitorquis (Quel.) Saccardo
= Agaricus rodmanii Peck
= Agaricus campestris var. edulis Vitt.

= Agaricus edulis (Vitt.) Moller and Schaeff.

Figure 144 Linear (longitudinally radial) mycelium of Agaricus bitorquis.

STRAINS: Horst B30 (The first commercial strain to be developed by Gerda Fritsche at the Dutch
Mushroom Research Center in Horst, Holland).

Horst K26, K32 (These are two second generation strains from Horst B30 and are distinctive
from it in that they fruit earlier, give higher yields and have slightly longer stems. Spawn of this
species is now available from Amycel.)

COMMON NAME: Rodman’s Agaricus

GREEK ROOT: Agaricus comes from the greek word “agarikon” which scholars believed origi-
nated with a Scythian people called Agari who were well versed in the use of medicinal plants and
employed a fungus called “agaricum”, probably a polypore in the genus Fomes. The species epi-
thet bitorquis means having two rings, for the double annulus that so distinguishes this species from
close relatives like Agaricus campestris, the Meadow Mushroom.

162/The Mushroom Cultivator

—

GENERAL DESCRIPTION: Cap smooth, white, thick fleshed, convex to broadly convex to plane
with age. The cap margin is incurved at first but soon decurves. The gills are pinkish at first, soon
darkening to chocolate brown with spore maturity. The stem is thick, relatively short and adorned
with a double membranous annulus. (The lower ring is often a thin annular zone). Its spores are
dark chocolate brown in mass.

NATURAL HABITAT: Naturally found in lawns, gardens, roadside areas, pastures, in enriched
grounds and on hard packed soil. A temperate species, widely distributed, A. bitorquis fruits pri-
marily in the spring and to a lesser degree in the fall.

GROWTH PARAMETERS

Mycelial Types: Rhizomorphic to linear; whitish to pale whitish in color.

Spawn Medium: Rye grain buffered with calcium carbonate and/or calcium sulfate. See Chapter
III.

Fruiting Substrate: Nitrogen enriched wheat straw and/or horse manure based compost bal-
anced to 71-74% moisture content.

Method of Preparation: See Chapter V on compost preparation. Pasteurization achieved
through exposure to live steam for 2 hours at 140° F. throughout the substrate. Compost should
be filled to a depth of 6-12 inches.

Spawn Run:

Relative Humidity: 90-100%.

Substrate Temperature: 84-86 °F. Thermal death limits have been established at 93 ° F. over
prolonged period of time.

Duration: 2 weeks.

C0 2 : 5,000-10,000 ppm.

Fresh Air Exchanges: 0 per hour.

Type of Casing: After fully run, cover with the standard casing whose preparation is described in
Chapter VIII. Layer to a depth of 1 -2 inches. The casing should be balanced to a pH of 7.2-7. 5.

Post Casing/Prepinning:

Relative Humidity: 90-1 00%.

Bed Temperature: 84-86 °F.

Duration of Case Run: 10-12 days.

C0 2 : 5000-10,000 ppm.

Fresh Air Exchanges: 0 per hour.

Primordia Formation:

Relative Humidity: 95-1 00%.

Bed Temperature: 77-80 °F.

Air Temperature: 15-71°F.

Growing Parameters for Various Mushroom Species/ 1 63

Lighting: None required.

C0 2 : less than 2000 ppm.

Fresh Air Exchanges: 2-4 per hour.

Watering: Regular misting (once to twice daily) of the beds stimulates primordia formation.

Cropping:

Relative Humidify: 85-92%.

Air Temperature: 75-77 °F.

C0 2 : less than 3000 ppm.

Fresh Air Exchanges: 2-4 per hour.

Flushing Interval: Every 8-9 days.

Harvest Stage: Directly before the partial veil stretches.

Yield Potential: Average commercial yields are reported at 3 Ibs/sq.ft. over a 5 week cropping
period. Maximum yields are 4 lbs per square foot.

Moisture Content of Mushrooms: 92% water; 8% dry matter.

Nutritional Content: Thought to be similar to Agaricus brunnescens.

Comments: The development of Agaricus bitorquis has given commercial growers greater flexi-
bility, especially those in warmer climates where elevated temperatures have been a limiting factor.
An advantage of this mushroom is its resistance to virus (a devastating disease that attacks A.
brunnescens) and its tolerance of high C0 2 levels. A disadvantage of growing this warmth-loving
Agaricus is the higher incidence of disease endemic to the temperature range in which this species
flourishes. Agaricus bitorquis is coarser, firmer, more strongly flavored and has a longer shelf life
than its close relative, A. brunnescens.

Genetic Characteristics: Basidia tetrapolar (4-spored), forming haploid spores, heterothallic. The
matinq of compatible monokaryons can result in fruitinq strains. Clamp connections absent. See
Chapter XV.

For further information consult:

P.J.C. Vedder 1978, “Modern Mushroom Crowing”, Educaboek, Culemborg, Netherlands.
(English edition available from Swiss American Spawn Company, Inc., Madisonville, Texas.)

P.J.C. Vedder 1 978, “The Cultivation of Agaricus bitorquis”] n The Biology and Cultivation
of Edible Mushrooms ed. by Chang and Hayes. Academic Press, New York.

Darmycel LTD. Spawn Lab Bulletin 1978, “A Guide to Darlington and Somycel Spawn
Strains”.

164/The Mushroom Cultivator

SPECIES: Agaricus brunnescens Peck

= Agaricus bisporus (Lge.) Sing.

Figure 145 Agaricus brunnescens fruiting in trays of compost.

STRAINS: Type or Brown Variety (var. bisporus)
White Variety (var. albidus)

Cream Variety (var. avellaneous)

COMMON NAME: The Button Mushroom.

GREEK ROOT: Agaricus comes from the greek word “agarikon” which scholars believed origi-
nated with a Scythian people called Agari who were well versed in the use of medicinal plants and
employed a fungus called “agaricum”, probably a polypore in the genus Fomes. The species epi-
thet brunnescens comes from the latin “brunneus” or brown. Literally, the name means the fungus
that becomes brown, probably referring to the color change of the flesh upon bruising. Also called
Agaricus bisporus for the two spored basidia populating the gill faces.

GENERAL DESCRIPTION: A robust, thick fleshed Agaricus species, with thin gills that are pink-
ish when young, and darkening to sepia and then chocolate brown in age. The cap is characteristi-
cally brownish, whitish or cream colored. The cap surface is smooth to appressed squamulose and
dry. This species has a short, thick stem which is adorned with a persistent membranous annulus
from a well developed partial veil. Its spores are chocolate brown in mass.

Crowing Parameters for Various Mushroom Species/ 1 65

NATURAL HABITAT: Naturally found in soils enriched with dung, on compost piles and in horse
stables. A temperate species, widely distributed, A. brunnescens fruits from May until November
over much of the northern hemisphere outside the tropical zone.

GROWTH PARAMETERS

Mycelial Types: Moderately rhizomorphic; dingy white, sometimes with brownish hues.

Spawn Medium: Rye grain buffered with calcium carbonate and/or calcium sulfate. See Chapter
III.

Fruiting Substrate: Nitrogen enriched wheat straw and/or horse manure based compost bal-
anced to 71-74% moisture content. This species also fruits well on rye grain covered with an un-
sterilized peat based casing layer.

Method of Preparation: See Chapter V on compost preparation. Pasteurization achieved
through exposure to live steam for 2 hours at 1 40 °F. throughout the substrate. Compost should be
filled to a depth of 6-12 inches.

1 66/The Mushroom Cultivator

Spawn Run:

Relative Humidity: 90-100%.

Substrate Temperature: 76-78 °F. Thermal death limits have been established at 96°F. but
damage can occur as low as 90 °F.

Duration: 2 weeks.

C0 2 : 5000-10,000 ppm.

Fresh Air Exchanges: 0 per hour.

Type of Casing: After fully run, cover with the standard casing whose preparation is described in
Chapter VIII. Layer to a depth of 1-2 inches. The casing should be balanced to a pH of 7. 0-7. 5.

Post Casing/Prepinning:

Relative Humidity: 90-100%.

Bed Temperature: 76-80 °F.

Duration of Case Run: 8-12 days.

C0 2 : 5000-10,000 ppm.

Fresh Air Exchanges: 0 per hour.

Primordia Formation:

Relative Humidity: 95-1007o.

Compost Temperature: 65-70 °F.

Air Temperature: 62-65 °F.

C0 2 : less than 1000 ppm.

Fresh Air Exchanges: 4 per hour.

Light: None required.

Cropping:

Relative Humidity: 85-92%.

Air Temperature: 62-65 °F.

C0 2 : less than 1000 ppm.

Fresh Air Exchanges: 4 per hour.

Flushing Interval: 7-10 days.

Harvest Stage: Directly before the partial veil stretches.

Light: None required.

Yield Potential: Average commercial yields are 3 lbs/ sq.ft, over a 5 week cropping period. Maxi-
mum yield is 6 lbs. per square foot.

Moisture Content of Mushrooms: 92% water; 8% dry matter.

Nutritional Content: 24-44' % protein (dry weight); 56 milligrams of niacin per 100 grams dry
weight.

Growing Parameters for Various Mushroom Species/ 1 67

Comments: Historically, this species and/or its close relatives were the first mushrooms to be culti-
vated in Europe during the late 1700’s. It remains the most widely cultivated mushroom in the
world today. A broad range of commercially available strains exist, many of which have been geneti-
cally selected for certain advantageous characteristics, especially yield, color and stature.

This species does not form pinheads on agar media unless activated charcoal or select bacteria
are present. A species sensitive to high levels of carbon dioxide, Agaricus brunnescens fruits only
within narrow environmental parameters. As a secondary decomposer, this species fruits best on
substrates that have been transformed by a succession of specific microorganisms.

The common button mushroom is the mainstay of the mushroom growing industry in this
country.

Genetic Characteristics: Basidia bipolar (2-spored), forming diploid spores; secondarily homo-
thallic. The mating of compatible dikaryons typically results in strains both more vigorous and high-
er yielding. Clamp connections absent. See Chapter XV.

For further information consult:

P.J.C. Vedder 1978, “Modem Mushroom Crowing”, Educaboek, Culemborg, Netherlands.
(English edition available from Swiss American Spawn Company, Inc., Madisonville, Texas).

Fred Atkins, 1973, “Mushroom Crowing Today”, MacMillan Publishing Co., New York.

SPECIES: Coprinus comatus (Mull, ex Fr.) Gray

168/The Mushroom Cultivator

STRAINS: On deposit at the American Type Culture Collection and available through various cul-
ture banks, both commercial and private.

COMMON NAME: The Shaggy Mane.

GREEK AND LATIN ROOTS: Coprinus comes from the Greek word “kopros” meaning dung
and comatus from the Latin “coma” meaning shaggy or adorned with hair tufts. The genus
Coprinus is noted for the several species that grow on dung and for deliquescing gills. The species
epithet describes the shaggy texture of the cap’s surface.

GENERAL DESCRIPTION: Cap medium to large in size, whitish, ovoid when young, soon elon-
gating upwards and becoming parabolic. As the mushroom matures and spores are produced, the
cap begins to disintegrate from the margin’s edge by an autodigestive process known as deliques-
cence. The disintegrating portions progressively darken and eventually liquify. The cap surface is

Figure 146 Fully mature Coprinus comatus fruiting in a tray of compost.

Growing Parameters for Various Mushroom Species/ 169

Figures 147-150 Four day developmental sequence of Coprinus comatus fruiting in a
tray of compost.

170/The Mushroom Cultivator

smooth at the disc, scaly below, soon gray, darkening with maturity until black and thin fleshed. The
gills are very crowded, whitish at first, soon gray, darkening with age to black. The partial veil mem-
branous, often leaving a fugacious, membranous (collar-like) annulus that can be moved over the
stem. The spore deposit is black.

NATURAL HABITAT: Common along roadsides, near debris piles, in lawns and in barnyards
during the late summer and fall.

GROWTH PARAMETERS

Mycelial Types: Linear to cottony, zonate-cottony mycelia; whitish in color.

Spawn Medium: Rye grain. See Chapter III.

Fruiting Substrate: Composted wheat straw enriched with horse and/or chicken manure, ad-
justed to 70% moisture content. Also, pasteurized chopped wheat straw supports fruitings of this
species. Garcha et al. (1979) reported that composts having the distinct scent of ammonia after
Phase II supported the greatest fruitings of Coprinus comatus.

Method of Preparation: See Chapters V & VI on the preparation of compost and straw. Pasteuri-
zation achieved through exposure to live steam for 2 hours at 140°F. Compost or straw should be
filled to a depth of 6-12 inches.

Spawn Run:

Relative Humidity: 90-100%.

Substrate Temperature: 76-80 °F.

Duration: 8-12 days.

C0 2 : 5000-10,000 ppm.

Fresh Air Exchanges: 0-1 per hour.

Type of Casing: After fully run, cover with the standard casing whose preparation is described in
Chapter VIII. Layer to a depth of 1-2 inches. The casing should be balanced to a pH of 7. 0-7. 5.

Post Casing/Prepinning:

Relative Humidity: 90-100%.

Bed Temperature: 76-80 °F.

Duration of Case Run: 10-12 days.

C0 2 : 5000-10,000 ppm.

Fresh Air Exchanges: 0-1 per hour.

Primordia Formation:

Relative Humidity: 95-100%.

Bed Temperature: 65-67 °F.

Air Temperature: 62-65 °F.

C0 2 : less than 1000 ppm.

Growing Parameters for Various Mushroom Species/ 171

Fresh Air Exchanges: 4 per hour.

Light: Natural daylight or grow-light recommended on a 1 2 hour on/ off cycle.

Cropping:

Relative Humidity: 85-92%.

Air Temperature: 62-65 °F.

C0 2 '. less than 1000 ppm.

Fresh Air Exchanges: 4 per hour.

Flushing Interval: 7 - 10 days.

Harvest Stage: Directly before the gills begin to deliquesce.

Light: Natural daylight or grow-light on a 12 hour cycle on/off cycle.

Yield Potential: Average commercial yields are 2-3 Ibs/sq.ft. over a 4 week cropping period.
Maximum yield potential has not yet been established.

Moisture Content of Mushrooms: 92-94% water; 6-8% dry matter.

Nutritional Content: 25.4 % protein (dry weight).

Comments: Like many other species in this genus, Coprinus comatus is a. thermotolerant
mesophile that often appears in compost piles. This mushroom was first grown in quantity at the
Dutch Mushroom Research Station using the same compost, casing and environmental parameters
as for the cultivation of Agaricus brunnescens. The authors have grown this species on compost
prepared for Agaricus and on straw alone, although fruitings appear more substantial on the former.

Coprinus comatus is edible and choice. However, the crops are difficult to keep because of the
early onset of deliquescence. By submerging mushrooms in water, deliquescence is slowed and
mushrooms remain in good condition for several days after picking.

Extracts from fresh specimens of this species has been shown to have antibiotic properties,
similar to those from Lentinus edodes.

Genetic Characteristics: Basidia tetrapolar (4-spored), forming haploid spores; heterothallic.
Clamp connections present. See Chapter XV.

For further information consult:

P.J.C. Vedder 1978, “Modern Mushroom Crowing”, Educaboek, Culemborg, Netherlands.
(English edition available from Swiss American Spawn Company, Inc., Madisonville, Texas).

172/The Mushroom Cultivator

SPECIES: Flammulina velutipes (Curt, ex Fr.) Sing.

= Collybia velutipes (Curt, ex Fr.) Kumm.

Figure 151 Flammulina velutipes fruiting in tray.

STRAINS: Many wild and domesticated strains of F. velutipes are available from commercial and
private stocks. (See Appendix). The Japanese have remained at the forefront of Enoke cultivation
with two popular commercial strains, “Maruei” and “Ebios”.

COMMON NAME: Enoke; Winter Mushroom; or Velvet Stem.

LATIN ROOT: Flammulina comes from the latin word “flammeus” or flame colored for the yel-
lowish orange to reddish orange color of the cap. The species epithet velutipes is the conjunction of
two latin words, the adjective “velutinus” meaning covered with fine hairs and the noun “pes” or
foot.

GENERAL DESCRIPTION: Caps typically small, reddish orange to reddish brown, at first hemis-
pherical, soon plane. The cap margin is often irregularly shaped. The gills are yellowish tinged. In
wild collections, the stem is densely fibri Hose, velvety, short and tough, in culture, however, the
stems are long and smooth. A partial veil is absent, its spores are whitish in mass.

Growing Parameters for Various Mushroom Species/ 173

NATURAL HABITAT: Common across the North American continent and in other temperate to
boreal regions of the world. Thriving on woody tissue, especially living trees and considered a cold
weather mushroom.

GROWTH PARAMETERS

Mycelial Types: Linear to cottony mycelia, sometimes aerial.

Spawn Medium: Sawdust/bran. One liter (1000 ml.) bottle of spawn inoculates 50-160 (800
ml.) containers. See Chapter III.

Fruiting Substrate: A 80-90% hardwood sawdust and 10-20% rice bran medium. Newly
chipped sawdust holds moisture poorly and some Japanese growers age the sawdust for several
years before using. Standard fruiting containers are quart mason jars or 800 ml. small mouthed
plastic bottles. Some growers are currently experimenting with the cultivation of this species on bulk
substrates in trays. Adjust moisture content of substrate to 58-60%.

Method of Preparation: See Chapter III for the preparation of sawdust/bran media. A 4:1 volu-
metric ratio of sawdust to bran (equivalent to a mass ratio of 1 0:1 sawdust to bran) is recommended.
Sterilize for 1-2 hours at 250 °F. (15 psi).

Spawn Run:

Relative Humidity: 90-1 00%).

Substrate Temperature: 72-77 °F.

Duration: 20-30 days using standard methods; 12-13 days using in vitro inoculation methods.
C0 2 : 5000-10,000 ppm.

Fresh Air Exchanges: 0 per hour.

Type of Casing: None required.

Primordia Formation:

Relative Humidity: 85%.

Air Temperature: 50-55 °F.

C0 2 : less than 1000 ppm.

Fresh Air Exchanges: 4 per hour.

Light: None needed.

Cropping:

Relative Humidity: 85%

Air Temperature: 50-55 °F.

Duration: 2-3 weeks.

C0 2 : less than 1000 ppm.

Fresh Air Exchanges: 4 per hour.

Light: Natural daylight or grow-light on a 1 2 hour cycle on/off cycle.

Flushing Interval: 1 0 days.

174/The Mushroom Cultivator

Figure 152 Developing pinheads of Flammulina velutipes.

Yield Potential: Average commercial yields are 160-220 grams per 800 ml. bottle. Maximum
yields are nearly 600 grams per 800 ml. bottle,

Moisture Content of Mushrooms: 92% water; 8% dry matter.

Nutritional Content: Reports vary from 1 8% to 3 1 % protein (dry weight); 1 07 milligrams of nia-
cin per 100 grams dry weight. F, Zadrazil (1979) found that colonization of straw by this species
decreases its digestibility for use as fodder. This contrasts with the effects of Pleurotus and
Stropharia rugoso-annulata whose presence on straw markedly increases its digestibility. Like many
wood degrading fungi, an anti-tumor antibiotic has been isolated from F. velutipes and is appropri-
ately called flammulin.

Comments: F. velutipes tends to form mycelial “pellets” soon after colonizing a substrate. This
phenomenon makes liquid culture techniques more difficult. Japanese researchers found that the
addition of 5% corn starch and 2% malt to a liquid solution inhibits the formation of these trouble-
some pellets. Curiously, fruitings on sawdust/bran beds can be precipitated when pieces of a fruit-
body are added to this solution. Shiio et al. (1974) found that one could induce the early formation

Growing Parameters for Various Mushroom Species/ 175

Figure 153 Flammulina velutipes fruiting in plastic
jar.

of fruitbodies with a technique whereby fresh pieces of Flammulina velutipes are mixed directly into
liquid spawn and then introduced into the sawdust/bran medium. Not only was the fruiting process
accelerated, the spawning period was cut in half and yield was nearly quadrupled over a year’s time.
Using this same technique with Pleurotus ostreatus, yields were increased over and above the norm
by a factor of three. Total production in either case, equalled as much as !4 of the substrate on a dry
weight basis. An analogous technique was developed by Urayama (1972) who discovered that cell-
free extracts of fresh F. velutipes mushrooms introduced to cultures of distantly related species
caused fruitbody formation.

Genetic Characteristics: Basidia tetrapoiar (4-spored), forming haploid spores; bifactorially het-
erothallic. Single spore isolates capable of producing sterile fruitbodies. Dikaryons are faster grow-
ing and characterized by clamp connections. Mycelium can produce oidia, self sectioning chains of
cells with similar functions as spores. See Chapter XV.

For further information consult:

H. Tonomura, 1974. “Flammulina velutipes’’ in The Biology and Cultivation of Edible
Mushrooms. Academic Press, New York.

176/The Mushroom Cultivator

Plate 1 Psilocybe cubensis mycelium. Plate 2 Psilocybe mexicana mycelium.

Plate 3 Lepista nuda mycelium. Plate 4 Psilocybe tampensis mycelium.

Plate 7 Compost raw materials at filling. Plate 8 Compost ready for spawning.

Note whitish colonies of Actinomycetes.

Plate 9 Agaricus brunnescens fruiting on cased horse manure compost.

Plate 10 Pleurotus ostreatus fruiting on pasteurized wheat straw.

Plate 1 1 Lentinus edodes, the
Shiitake mushroom, fruiting on
oak logs.

Plate 12 Coprinus comatus,
the Shaggy Mane, fruiting on
cased horse manure compost.

Plate 1 3

Psilocybe mexicana,
Teonanacatl, fruiting
on cased rye grass
seed.

Plate 1 4 Psilocybe
tampanensis sclerotia
on rye grass seed.

Plate 15 Psilocybe
tampanensis fruiting
on cased rye grass
seed.

SPv e- '®»ra!w&» , 5£sSS i a»

Plate 16 Psilocybe cyanescens mycelium running through moist alder sawdust.

Plate 18 Psilocybe cubensis fruiting on cased, pasteurized wheat straw

Plate 20 Penicillium, the Blue Green Plate 21 Aspergillus, the Green Mold
Mold and Cladosporium, the Dark Green growing on malt agar media.

Mold, growing on malt agar media.

Growing Parameters for Various Mushroom Species/ 1 77

STRAINS: Numerous strains of Lentinus edodes are available from commercial and private
stocks. The American Type Culture Collection, which sells cultures to educational organizations
and research facilities, has stock cultures of several wild and domesticated strains. Strains are often
distinguished by their preferences for fruiting in colder or warmer temperature zones.

COMMON NAMES: The Shiitake Mushroom; The Japanese Black Mushroom; and The Chinese
Black Mushroom. (The name shiitake comes from the association of this mushroom to the shiia
tree, a member of the genus Pasania).

LATIN AND GREEK ROOTS: Lentinus comes from Mentis” or lens-shaped for the form of the
cap and edodes signifies the edibility of this species.

GENERAL DESCRIPTION: Cap pale to dark reddish brown, convex, becoming broadly convex
to nearly plane in age. The cap margin is typically in rolled when young. The cap surface is covered
with whitish veil remnants, especially along the margin. The flesh is firm, pliant, easily drying and re-
constituting. The gills are whitish, close to crowded, often with serrated edges. The stem is centrally
attached to the cap, short, very tough and adorned with scattered fibrillose remnants of the partial
veil. Its spores are whitish in mass.

NATURAL HABITAT: A wood decomposer, typically saprophytic. Lentinus species are common
on the dead tissue of deciduous trees, mainly Fagaceae (oak, chestnut, shiia [Pasania] and beech).
In nature, they particularly prefer oaks. Fruiting in the fall, early winter and spring, this species is in-
digenous to Japan, China and other countries in the temperate zone of the Indo-China region.

GROWTH PARAMETERS

Mycelial Types: Rhizomorphic to linear.

Spawn Medium: Pre-soaked wooden dowels or a^kl sawdust/bran mixture. See Chapter III.

\ Zj

Fruiting Substrate and Method of PreparatiM:2|ak or alder logs, 4-6 inches in diameter, are
sawed into 3 foot lengths. These logs should bbjjay|sfie spring or fall to maximize sap content and

sawed into 3 foot lengths. These logs should bkcJs4Ji|jfhe spring or fall to maximize sap content and
can be inoculated immediately. (Some growers season their logs in shaded, open air stacks
for one month prior to inoculation). Before inoculating, logs should be cleaned of any lichen or
fungal growths.

Alternative fruiting substrates include alder or oak sawdust and bran mixed 4:1 with a moisture
content of 60% and sterilized at 1 5 psi for 1 -1 Vz hours. Fortified rye grass straw has also been used
as a sterile fruiting medium. (See Chapter III).

Spawn Run:

Relative Humidity: 60-75% for logs; 90% for sawdust.

Substrate Temperature: Fast growth at 77 °F. (Temperatures above 95 °F. and below 41 °F.
stop mycelial growth).

Duration: 6-12 months for cut logs; 30-60 days for sawdust blocks.

C0 2 : None established; no controls needed using these methods.

178/The Mushroom Cultivator

Fresh Air Exchanges: Stacks in open air sufficient. (Recent innovations show that logs stacked
in a vertical configuration and covered with straw and plastic to maintain even temperatures
result in faster spawn running in an outdoor environment. Within a controlled greenhouse, the
logs need not be covered. The contact between the log surfaces should be minimized to pre-
vent competitor molds and lichens from forming).
pH Optima: 5-6.

Light: None required.

Type of Casing: None needed.

Pinhead Initiation:

Initiation Technique: Submerge logs and blocks in cold water for 24-72 hours.

Relative Humidity: 95%.

Air Temperature: 59-68 °F.

Duration: 7-14 days after soaking.

C0 2 : Not applicable.

Fresh Air Exchanges: If within a greenhouse, 2-4 per hour.

Light: Ambient natural light or optimally 10 lux in the 370-420 nanometer range.

Cropping:

Relative Humidity: 85-90%.

Air Temperature: 59-68 °F.

C0 2 : less than 1000 ppm.

Fresh Air Exchanges: 2-4 per hour or sufficient to meet C0 2 and/or cooling requirements.
Duration: 3-5 years on oak logs; 2-3 years on alder.

Harvest Stage: Directly before the incurved margin straightens and the cap expands to plane.
Flushing Interval: Outdoor methods generate 2 flushes per year (fall and spring); indoor meth-
ods can produce up to 4 flushes depending on the soaking/initiation schedule.

Light: Same as above.

Yield Potential: Average commercial yields are 2-3 lbs (fresh weight) of mushrooms per log.
Moisture Content of Mushrooms: 85% water; 15% dry matter.

Nutritional Content: 1 0.0-1 7.5% crude protein (dry weight) and 55 milligrams of niacin per 1 00
grams dry weight.

Comments: Compounds in this mushroom have anti-cholesterol effects. Chihara (1979) reported
that lentinan, a water soluble polysacharide in L. edodes, was “found to almost completely regress
the solid type tumors of sarcoma-180 and several kind (sic) of tumors. . . The work of others
(Cochran, 1 978; Tokita et al., 1 972; Tokuda and Kaneda, 1 979) have similarly described the ben-
eficial properties of this fungus. (See Appendix III).

Although the standard method of cultivation involves oak logs, recent experiments employing

Growing Parameters for Various Mushroom Species/ 1 79

sawdust or rye grass based “synthetic” mixtures have proved that Lentinus edodes can be grown
on a variety of substrates. In a recent article, Han et alia (1981) report the results of growth experi-
ments with shiitake mini-logs composed of 90% broadleaf sawdust, 10% rice bran and 0.2%
CaC0 3 . Supplements that increased mycelial growth more than rice bran were yeast powder
(2.0%), soybean meal (5.0%), milk powder (2.0%) and molasses (1.5%). The fastest mycelial
growth occurred when the moisture content of the logs was balanced to 50-60%. In tests on fruiting
and yield the following data were compiled:

Once mycelial growth is complete, highest yields were achieved if the vegetative cycle was pro-
longed 4-12 weeks, with the maximum yield at 12 weeks.

At pin initiation, water bath periods of 48-72 hours increased the moisture content of the logs
by 5-15% and yields by 50%.

Cooling the logs for eight days at 60-62 °F. following 48 hours of soaking gave the highest
yields.

Using 0. 1 % IN hydrochloride to adjust the pH of the water bath from 4. 5-7.0, a pH of 5.0 pro-
duced the most primorida and mature mushrooms.

At a light intensity of 550 lux, yields were highest.

The addition of the hormones NAA (5ppm), gibberellin (lOppm), ethylene chlorohydrin
(2000x) and colchicine (8000x) as well as yeast powder (0.1 %) to the water bath increased
yields.

Nevertheless, the traditional log method remains the most commercially feasible at this time
and the one best suited to home cultivation.

Genetic Characteristics: Basidia tetrapolar, forming four haploid spores; heterothallic. Dikaryons
with clamp connections. See Chapter XV.

For more information consult:

H. Akiyama et al., 1974. “The Cultivation of Shii-ta-ke in a Short Period”. Mushroom
Science IX, pp. 423-433.

T. Ito, 1978. “Cultivation of Lentinus edodes” in The Biology and Cultivation of Edible
Mushrooms Ed. by S.T. Chang, pp. 461-473.

R. Kerrigan, 1982. “Is Shiitake Farming for You?” Far West Fungi, Santa Cruz.

Y.H. Han, W.T. Veng and S. Cheng, 1981. “Physiology and Ecology of Lentinus edodes
(Berk.) Sing.” Mushroom Science XI, Melbourne.

SPECIES: Lepista nuda (Bull, ex Fr.) Cooke

= Clitocybe nuda (Fr.) Bigelow and Smith
= Tricholoma nudum (Fr.) Kummer

Figure 155 Mycelium of Lepista nuda.

STRAINS: Available from commercial and private stocks. The American Type Culture Collection
has several strains. Although few spawn companies sell strains of L. nuda, tissue and spore cultures
are easily obtained from wild specimens. Nevertheless, there are a limited number of productive
strains currently in circulation.

COMMON NAME: The Blewit.

LATIN AND GREEK ROOTS: Lepista comes from the greek “lepis” which means scale. On the
other hand, the species epithet nuda comes from “nudus” or naked. The name Lepista nuda com
stitutes a contradition of terms, literally translating as the scaly smooth mushroom.

GENERAL DESCRIPTION: Cap typically violet when fresh, becoming buff brown in drying;
smooth, without hairs; dry; convex or broadly convex to plane in age. The cap margin is inrolled or
incurved when young and simply decurved at maturity. The gills are a pale violet color, sometimes
developing brownish hues in age and are adnexed or ascending in their attachment to the stem. The
stem is equal overall but bulbous at the base and covered with fine fibrils over much of its surface.

Growing Parameters for Various Mushroom Species/ 181

Fruitbodies can be moderately large when mature. A partial veil is absent. The spore deposit is pale
pinkish tan.

NATURAL HABITAT: Commonly occurring in the summer to late fall across much of the tem-
perate regions of North America and Europe. This species is found in and around decomposing
piles of sawdust, in conifer duff, amongst leaves and in mature compost piles.

GROWTH PARAMETERS

Mycelial Types: Linear to cottony and usually with purplish to violet hues. (See Color Photo 3).
Spawn Medium: A 4:1 sawdust/bran mixture or rye grain spawn. See Chapter III.

Fruiting Substrates: Horse manure/straw compost mixed with 1 0% fresh straw at spawning; leaf
mulch/sawdust mixtures.

Spawn Run:

Relative Humidity: 90 + %.

Substrate Temperature: Fastest growth at 70-75°F. Temperature maxima and minima: 40°F.
and 86 °F. respectively.

Duration: 25-60 days for complete colonization.

C0 2 : 5000-10,000 ppm.

Fresh Air Exchanges: 0 per hour.

Light: Incubation in total darkness.

Type of Casing: Standard peat based casing. An option is the addition of shredded leaf material
and activated charcoal to 10% of total mass. Balance to a pH of 7.0.

Pinhead Initiation:

Relative Humidity: 95%.

Air Temperature: 55-65 °F.

Duration: 7-14 days.

C0 2 : less than 1000 ppm
Fresh Air Exchanges: 2-4 per hour.

Light: Ambient natural light or optimally 10 lux in the 370-420 nanometer range. (Light re-
quirements have not yet been established for this species, and until that time, light stimulation
should be presumed as a prerequisite for fruiting.)

Cropping:

Relative Humidity: 85-90%.

Air Temperature: 55-65 °F.

Duration: 24-52 weeks.

C0 2 : less than 1000 ppm.

Fresh Air Exchanges: 2-4 per hour.

182/The Mushroom Cultivator

FVli' J fflhyif r 'lT -tf i - , ;nj —

Harvest Stage: While the mushroom caps remain convex.

Flushing Interval: 10-14 days.

Light: Same as above.

Yield Potential: Data very limited. Yields of one and a quarter pounds per square foot in 1 4 weeks
have been reported by Visscher (1981). (Recent studies show that yields can be increased substan-
tially, although no maxima have yet been established.)

Moisture Content of Mushrooms: 88-90% water; 10-12% dry matter.

Nutritional Content: No data available.

Comments: Several contraditions about the fruiting requirements for this species are appar-
ent. Although Wright and Hayes (1979) reported that immature horse manure/straw composts
supported the most vigorous mycelial growth, the work of previous researchers indicates that the
best fruitings occurred on “spent” compost that has been colonized for a year or more. Fruitbodies
also form on spawned leaf mulch mixed with sawdust. The fruiting mechanism may, in part, be con-
trolled by bacterial flora associated with leaf mulch and the decomposition process.

Singer (1963) reported that mycelium implanted in beds of horse manure/straw compost for
7-1 4 months produced mushrooms directly after the appearance of rhizomorphs. J. Garbaye et al.
(1 979) published data indicating that the supplementation of natural patches with a NPKCa mineral
fertilization induced large fruitings of L. nuda as well as Boletus edulis and Lepiota rachodes, two
unrelated species of culinary distinction.

Alexander Smith (1980) remarks that this mushroom should not be eaten raw, but only after
cooking. European books have reported that this mushroom contains thermobile hemolysin, a
compound that degenerates red blood cells. Although this mushroom has been responsible for scat-
tered poisonings when quantities have been eaten, the effects have been relatively minor and the
toxin is easily destroyed by cooking or parboiling. Lepista nuda is, however, a mushroom with
many positive attributes. Its striking color, firm texture and good taste recommend this species as
one of high culinary appeal.

Some commercial production of L. nuda is ongoing in Europe. Nevertheless, this mushroom
is not, as of yet, a species with yields substantial enough to warrant commercial production in this
country. It is a mushroom more suited to the interests of home cultivators and natural culture tech-
niques.

Genetic Characteristics: Basidia tetrapolar, forming four haploid spores; heterothal lie. Dikaryons
with clamp connections. See Chapter XV.

For more information consult:

S.H. Wright and W.A. Hayes, 1979. “Nutrition and Fruitbody Formation of Lepista Nuda
(Bull, ex Fr.) Cooke”, pp. 873-884 in Mushroom Science X, Part I. Bordeaux.

J. Garbaye et alia, 1979. “Production De Champignons Comestibles En Foret Par Fertilisa-
tion Minerale-Premiers Resultats Sur Rhodopaxillus Nudus”. pp.81 1 -81 6 in Mushroom Science
X, Part I. Bordeaux.

M. Vaandrager and H.R. Visscher, 1981. Experiments on the Cultivation of Lepista Nuda, the
Wood Blewit”, pp. 749-759 in Mushroom Science XI, Australia.

Growing Parameters for Various Mushroom Species/ 1 83

SPECIES: Panaeolus cyanescens Berkeley and Broome
= Copelandia cyanescens (Berk. & Br.) Sing.

Figure 156 Panaeolus cyanescens fruiting on cased straw.

184/The Mushroom Cultivator

nrz

STRAINS: Hawaiian.

Mexican.

COMMON NAME: Pan cyan.

GREEK ROOT: Panaeolus is Greek for “all variegated’’, in reference to the spotted appearance of
the gills. The species name cyanescens comes from “cyaneus” or blue for the color the flesh be-
comes upon bruising.

GENERAL DESCRIPTION: Cap 15-40 mm. broad. Hemispheric to campanulate to convex or
broadly convex at maturity. The margin is initially shortly translucent striate when wet, opaque when
dry. The cap is light brown at first, becoming pallid grey in drying, eventually pallid to white, often
covered with spores. The gills are adnexed in their attachment, close, thin, with two or three tiers of
intermediate gills and mottled grayish black at with spore maturity. The stem is 85-120 long X
1 5-30 mm. thick and equal to bulbous at the base, tubular, often grayish towards the apex, pale yel-
lowish overall, and flesh colored to light brown towards the base. The flesh readily turns bluish
where bruised. A partial veil is absent. Its spores are dark violet-black.

NATURAL HABITAT: Scattered to numerous on dung, in well manured grounds, grassy areas,
meadows, or pastures. Known from Hawaii and Mexico. Two other Panaeoli, close to P.
cyanescens macroscopically and microscopically, grow in western Washington and in Florida.

GROWTH PARAMETERS

Mycelial Types: Linear to cottony mycelia; white to off-white, sometimes bruising bluish where in-
jured.

Spawn Medium: Rye grain. See Chapter III.

Fruiting substrate: Pasteurized wheat straw; horse manure/straw compost.

Method of Preparation: Chopped wheat straw pasteurized in a hot water bath at 160° for 20-30
minutes, cooled and spawned or horse manure/ straw compost prepared according methods out-
lined in Chapter V.

Spawn Run:

Relative Humidity: 90 + %.

Substrate Temperature: 79-84° F.

Duration: 7-12 days.

C0 2 : 10,000 ppm or higher.

Fresh Air Exchanges: 0 per hour.

Type of Casing: Standard peat based casing whose preparation is described in Chapter VIII. Layer
to a depth of VzA inch.

Post Casing/Pre-pinning:

Relative Humidity: 90 + %.

Substrate Temperature: 79-84 °F.

Growing Parameters for Various Mushroom Species/ 1 85

C0 2 : 10,000 ppm or above.

Fresh Air Exchanges: 0 per hour.

Light: Incubation in darkness.

Primordia Formation:

Relative Humidity: 95 + %.

Air Temperature: 75-80° F.

C0 2 : 5,000 ppm or below.

Fresh Air Exchanges: 2 per hour.

Light requirements: Diffuse natural or fluorescent grow-lights.

Cropping:

Relative Humidity: 85-92%.

Air Temperature: 75-80° F.

C0 2 : 5,000 ppm or below.

Fresh Air Exchanges: 2 per hour.

Harvest Stage: When the caps are convex.

Flushing Interval: 5-7 days.

Light: Diffuse natural or grow-lights.

Yield Potential: Not yet established.

Moisture Content: 90-92% water; 8-10% dry matter.

Comments: This rapidly growing species fruits readily on pasteurized straw provided a thin layer of
casing is applied (Vz inch). No more than one week passes from the time of casing to the first flush.
Although the fruitbodies are small, the flushes are typically abundant. The degree of bluing seems to
vary with the strain and substrate.

186/The Mushroom Cultivator

SPECIES: Panaeolus subbalteatus Berkeley and Broome
= Panaeolus venenosus Morrill

Figure 157 Panaeolus subbalteatus fruiting
outdoors on horse manure- wood chip compost.

STRAINS: Fruiting strains are easily obtained from wild specimens.

COMMON NAME: The Belted Cap Panaeolus.

GREEK ROOT: Panaeolus is Greek for “all variegated” in reference to the spotted appearance of
the gills. The species name subbalteatus comes from the conjunction of the prefix “sub-” meaning
almost or somewhat and “balteatus” or belt-like, for the characteristic color zonation that forms
along the margin of the cap in drying,

— rr r

Growing Parameters for Various Mushroom Species/ 187

GENERAL DESCRIPTION: Cap 35-50 mm. broad at maturity. The cap is convex to campanu-
late, then broadly convex and finally expanding to nearly plane with a broad umbo. The color is cin-
namon brown to orangish cinnamon brown, fading to tan in drying with a dark brown encircling
zone along the cap margin. The gills are attached to the stem, broader at the center and with three
tiers of intermediate gills inserted. The gill color is brownish and spotted, with the edges remaining
whitish, becoming blackish overall from spore maturity. The stem is 50-60 mm. long by 4 mm.
thick at maturity and is brittle, hollow, fibrous, and enlarges towards the base. The color is reddish
toned beneath a fine sheath of minute whitish fibrils, darkening downwards or when touched. The
stem base often bruises bluish. On the cap, bluing is rarely seen.

NATURAL HABITAT: Scattered to numerous on stable leavings from horses; in horse dung; or
in well manured grounds. This species is widely distributed across the North American continent
and throughout temperate regions of the world.

GROWTH PARAMETERS

Mycelial Type: Cottony mycelia noted; whitish to off-white in color.

Spawn Medium: Rye grain.

Fruiting Substrate: Horse manure compost, pasteurized wheat straw.

Method of Preparation: Horse manure/straw compost or pasteurized wheat straw prepared ac-
cording to methods outlined in Chapters V & VI respectively.

Spawn Run:

Relative Humidity: 90 +%.

Substrate Temperature: 80-86 °F.

Duration: 7-12 days.

C0 2 : 10,000 ppm or higher.

Fresh Air Exchanges: 0 per hour.

Type of Casing: Casing optional. If used, make up a standard peat based casing whose prepara-
tion is described in Chapter VIII. Layer to a depth of Zi to 1 inch.

Post Casing/Pre-pinning:

Relative Humidify: 90%.

Substrate Temperature: 80-86 °F.

C0 2 : 10,000 ppm or above.

Fresh Air Exchanges: 0 per hour.

Light: Incubate in darkness.

Primordia Formation:

Relative Humidify: 95 + %.

Air Temperature: 75-80 °F.

C0 2 : 5,000 ppm or below.

188/The Mushroom Cultivator

Fresh Air Exchanges: 2 per hour.

Light: Diffuse natural or grow-lights.

Cropping

Relative Humidity: 85-92%.

Air Temperature: 75-80 °F.

C0 2 : 5,000 ppm or below.

Fresh Air Exchanges: 2 per hour.

Harvest Stage: When the caps have expanded to nearly plane.

Light: Diffuse natural or grow-lights.

Yield Potential: Not yet established.

Moisture Content: 90-927o water; 8-10% dry matter.

Comments: Panaeolus subbalteatus is a fast running and an early fruiting mushroom that easily
grows in controlled environments. Possessing low levels of psilocybin and/ or psilocin, the fruit-
bodies are small compared to other cultivated mushrooms. Hence, it has not been as popular with
home cultivators as for instance, Psilocybe cubensis.

Given the fact that Panaeolus cyanescens fruits well on pasteurized wheat straw, Panaeolus
subbalteatus is likely to fruit on that substrate as well. Pollock (1977) fruited this species on cased
crimped oat spawn. Undoubtedly, Panaeolus subbalteatus can be grown on a wide variety of sub-
strates.

Short term “natural culture” of this mushroom is also possible although yields are much lower
than those attained in a controlled indoor growing environment. Horse manure/ straw compost ar-
ranged in outdoor beds can be inoculated with mycelium from wild patches or grain spawn can be
used.

Panaeolus subbalteatus is considered a “weed mushroom” by commercial Agaricus growers
and its presence suggests under-composting and/ or excessive moisture. This species once had the
reputation, albeit undeserved, of being poisonous— thus the synonym Panaeolus venenosus.

Growing Parameters for Various Mushroom Species/ 189

SPECIES: Pleurotus ostreatus (Jacq. ex Fr.) Kummer

STRAINS: Strains of Pleurotus ostreatus are available from commercial and private stocks. The
American Type Culture Collection, which sells cultures to educational organizations and research
facilities, has stock cultures of several wild and domesticated strains. Somycel’s-3004 is the stan-
dard strain used by the European Pleurotus industry and is synonymous with ATCC’s-38546.

COMMON NAME: The Oyster Mushroom.

LATIN AND GREEK ROOTS: Pleurotus comes from the greek “pleuro” which means formed
laterally or in a sideways position, referring to the lateral position of the stem relative to the cap. The
species epithet ostreatus refers to its oyster shell-like appearance and color.

GENERAL DESCRIPTION: Cap tongue shaped, maturing to a shell shaped form, 50-1 50 mm.
in diameter; whitish to gray to blue gray overall. (Color is a light determined factor in this species).
The flesh is thin and white. The margin is even and occasionally wavy. The gills are white, decurrent

Figure 158

Fully mature Pleurotus ostreatus mushrooms fruiting on straw.

Figures 159-162 Four day developmental sequence of Pleurotus ostreatus fruiting on
wheat straw.

and broadly spaced. The stem is attached in an off-centered fashion and is short at first and absent in
age. Its spores are whitish to lilac gray in mass.

NATURAL HABITAT: A wood decomposing, saprophytic or parasitic fungus. Pleurotus
ostreatus grows abundantly on standing and fallen alder, cottonwood and maple. This species is es-
pecially numerous in river valleys and fruits in the fall, early winter and spring across much of tem-
perate North America.

Growing Parameters for Various Mushroom Species/ 191

GROWTH PARAMETERS

Mycelial Types: Fast growing rhizomorphic to linear mycelia noted. Color is typically whitish.
Spawn Medium: Rye grain. See Chapters III.

Fruiting Substrate and Method of Preparation: Cereal straw (normally wheat) balanced to a
75% moisture content. The straw, chopped or whole, is pasteurized by submerging in a 160°F.
water bath for 30-45 minutes. An alternative method utilizes live steam pasteurization at 140°F. for
6 hours. In Japan, Pleurotus is grown on a mixture of hardwood sawdust and bran (4 parts to 1 ,
65% moisture and a pH of 6. 8-7.0). This mixture is sterilized for 1-2 hours at 15 psi. Being a
primary decomposer. Pleurotus grows on a wide variety of cellulosic wastes.

Spawn Run:

Relative Humidity: 90-100%.

Substrate Temperature: Fastest growth at 78-84°F. Thermal death occurs if mycelium is held
above 104°F. for 48 hours.

Duration: 10-14 days for complete colonization.

C0 2 : 20,000 ppm or 20% C0 2 by volume. (Growth is stimulated up to 28,000 ppm).
Fresh Air Exchanges: 0 per hour.

Light: Incubation in total darkness.

Type of Casing: None needed.

Pinhead Initiation:

Relative Humidity: 95%.

Air Temperature: 55-60 °F.

Duration: 7-14 days.

C0 2 : less than 600 ppm.

Fresh Air Exchanges: 4 per hour.

Light: Phototropic, most responsive to an exposure of 2,000 lux/hour for 12 hours/ day.
Grow-lux type fluorescent lighting is recommended. Diffuse natural light is sufficient.

Watering: Regular misting once to twice daily until fruitbodies are 30-40% of harvest size and
then water as needed to prevent caps from cracking.

Cropping:

Relative Humidity: 85-92%.

Air Temperature: 60-64 °F.

Duration: 5-7 weeks.

C0 2 : less than 600 ppm.

Fresh Air Exchanges: 4-6 per hour or sufficient to meet C0 2 and/ or cooling requirements.
Harvest Stage: Directly before incurved margin elevates to plane.

Flushing Interval: 1 0 days.

Light: Same as above.

l^^The M us h room C u l t iv ator

Watering: Regular misting to prevent caps from cracking and to keep resting pinheads viable.
Yield Potential: Average commercial yields are 1 kilogram fresh weight of mushrooms per
kilogram of dry weight of straw substrate.

Moisture Content of Mushrooms: 91% water; 9% dry matter.

Nutritional Content: Crude protein has been reported at 30.4 % of dry weight and 109 milli-
grams of niacin per 100 grams dry weight.

Comments: Biologically, Pleurotus ostreatus efficiently utilizes its substrate. Its ability to fruit on a
single component substrate, to permeate the straw rapidly while tolerating high carbon dioxide
levels and to produce abundant crops within a short time period, make Pleurotus ideal for home
cultivation.

Of concern to cultivators growing in enclosed rooms is the abundant spore load generated by
this species. Pleurotus spores cause allergic reactions amongst some workers and mycophagists.
Sporeless strains are therefore desirable and are the object of current research. Eger (1974) noted
the possibility that heavy spore concentrations from Pleurotus farms could infect surrounding wood-
lands.

Pleurotus ostreatus var. Florida , a warmth loving relative, is also cultivated in Europe (Hungary)
and shares many of the growth properties of Pleurotus ostreatus.

Genetic Characteristics: Basidia tetrapolar, producing 4 haploid spores; heterothallic. Clamp
connections present. See Chapter XV.

For more information consult:

F. Zadrazil, 1 974. “The Ecology and Industrial Production of Pleurotus ostreatus, Pleurotus
Florida, Pleurotus cornucopiae, and Pleurotus eryngii” in Mushroom Science IX (Part I), The
Mushroom Research Institute, Japan.

Growing Parameters for Various Mushroom Species/ 1 93

SPECIES: Pleurotus ostreatus (Jacq. ex Fr.) Kummer (Florida

variety)

= Pleurotus ostreatus var. florida nom. prov. Eger
= Pleurotus floridanus Singer

STRAINS: Most strains of this mushroom originate from wild specimens cultivated in 1958 by
S.S. Block of Gainesville, Florida. Eger compared the Florida strains with Pleurotus ostreatus from
Michigan (supplied by Alexander Smith) and found them to be identical in form, taste, color and
odor. Spore size and shape are also the same. Monokaryons arising from single spore germinations
are completely cross fertile, suggesting that these two mushrooms are not separate species, but dif-
ferent strains within the same species.

The American Type Culture Collection, which sells cultures to educational organizations and
research facilities, lists this mushroom under Pleurotus ostreatus as number #38538. This strain is
Block’s original. Eger returned to Florida with San Antonio in 1977 and recollected four more
strains of Pleurotus, three of which were deposited with ATCC. They are respectively: FI = ATCC
#38539; F2 = #38540; F4 = #38541 .

The Florida Pleurotus is available as commercial spawn from Somycel as #3025. The Swiss
American Spawn Company sells a “low spore load” strain called P-3.

COMMON NAME: Pleurotus Florida. The Florida Pleurotus.

LATIN AND GREEK ROOTS: Pleurotus comes from the Greek “pleuro” which means formed
laterally or in a sideways position, referring to the lateral position of the stem relative to the cap. The
epithet Florida obviously refers to the locality where this mushroom was first collected.

GENERAL DESCRIPTION: Cap tongue shaped, maturing to a shell shaped form, 50-100 mm.
in diameter; whitish to gray to pale yellow brown. (Color is a light and temperature determined fac-
tor in this species). The flesh is thin and white. The margin is even and occasionally wavy. The gills
are white, decurrent and broadly spaced. The stem is attached in an off-centered fashion and is short
at first and absent in age. Its spores are whitish to lilac gray in mass.

GROWTH PARAMETERS

Mycelial Types: Fast growing rhizomorphic to linear mycelia. Its color is typically whitish.
Standard Spayvn Medium: Rye grain. See Chapter III.

Fruiting Substrate and Method of Preparation: Cereal straw (normally wheat) balanced to a
75% moisture content. The straw, chopped or whole, is pasteurized by submerging in a 160°F.

1 94/The Mushroom Cultivator

water bath for 20-30 minutes. An alternative method utilizes live steam pasteurization at 140°F. for

In Japan, Pleurotus is grown on a mixture of hardwood sawdust and bran (4 parts to 1 , 65%
moisture and’a pH of 6. 8-7.0). This mixture is sterilized for 1 hour at 15 psi. Being a primary de-
composer, Pleurotus grows on a wide variety of wastes high in cellulose.

Spawn Run:

Relative Humidity: 90-100%.

Substrate Temperature: Fastest growth at 82-86°F. Thermal death occurs if mycelium is held
above 104°F. for 72 hours.

Duration: 10-14 days for complete colonization.

C0 2 : 20,000 ppm or 207o C0 2 by volume. (Growth is stimulated up to 28,000 ppm).
Fresh Air Exchanges: 0 per hour.

Light: Incubation in total darkness.

Type of Casing: None needed.

Pinhead Initiation:

Relative Humidity: 95%.

Air Temperature: 72-77 °F.

Duration: 7-14 days.

C0 2 : less than 600 ppm
Fresh Air Exchanges: 4 per hour.

Light: Positive phototropism has been firmly established. 2,000 lux/hours for 1 2 hours/ day is
most stimulatory. Grow-lux type fluorescent lighting is recommended. Diffuse natural light is
sufficient.

Watering: Regular misting (once to twice daily) of the substrate until the fruitbodies are
30-40% of harvest size.

Cropping:

Relative Humidity: 85-92%.

Air Temperature: 72-77 °F.

Duration: 4-5 weeks.

C0 2 : less than 600 ppm.

Fresh Air Exchanges: 4-6 per hour or sufficient to meet C0 2 and/ or cooling requirements.
Harvest Stage: Directly before incurved margin expands to plane.

Flushing Intervals: 1 0 days.

Light: Same as above.

Watering: Misting recommended to prevent cracking of caps and to prevent resting primordia
from drying.

Yield Potential: Average commercial yields for Pleurotus ostreatus var. Florida are 1 kilogram

Growing Parameters for Various Mushroom Species/ 1 95

fresh weight of mushrooms per kilogram of dry weight of straw substrate. Pleurotus ostreatus var.
florida produces more mushrooms within a shorter period of time while attaining a similar total yield
per dry pound of substrate than does Pleurotus ostreatus.

Moisture Content of Mushrooms: 91% water; 9% dry matter.

Nutritional Content: Crude protein has been reported at 30.4% of dry weight and 109 milli-
grams of niacin per 1 00 grams dry weight.

Comments: The Floridan Pleurotus ostreatus, a warmth loving variety, is popular with growers in
Europe (Hungary, France and Germany) and shares many of the ggs»vth characteristics of Pleurotus
ostreatus. Its preference for warmer climes recommends this spadA for cultivation during the late
spring through early fall whereas P. ostreatus is ideal for uVrpJ(i!$tivation.

This mushroom, like its close cousin P. ostreatus, is peNmt ij.4 the home cultivator. But, the
Floridan Pleurotus ostreatus has a distinct advantage over P. ostreatus in that a “cold shock” is not
needed for pinhead formation and the period from initiation to first flush is only 1 0 days compared
to 20 days for P. ostreatus. Its ability to fruit on a singular substrate, to permeate the straw rapidly
while tolerating high C0 2 levels and to produce abundant crops within a short time frame, makes
Pleurotus an excellent species for small scale cultivation.

The taxonomy of this “species” is unsettled and contradictory. Dr. Rolf Singer places P.
floridanus in the Section Lentodiellum whose species are characterized by deeply rooted metuloid
pleurocystidia (sterile surface cells on the gill having incrustations) and have mycelia that do not
sclerotize. On the other hand, he assigns Pleurotus ostreatus to the type section Pleurotus which
lacks metuloid pleurocystidia and has hyphae that undergoes sclerotization. Since monokaryons
from single spores are compatible between these two mushrooms, and because sporulating fruit-
bodies form as a result of their mating, it seems clear that these two mushrooms are one species
sharing a common genetic heritage.

Of concern to cultivators is the abundant spore load produced by this mushroom, most notice-
able within an enclosed growing environment. Some people suffer allergic reactions when coming
into contact with Pleurotus spores. A small fraction of mycophagists are unable to eat P. ostreatus
and allies without stomach upset. Hence, when eating these mushrooms for the first time, small por-
tions are recommended.

Genetic Characteristics: Basidia tetrapolar, producing 4 haploid spores; heterothallic. Clamp
connections present. See Chapter XV.

For more information consult:

F. Zadrazil, 1 974. “The Ecology and Industrial Production of Pleurotus ostreatus, Pleurotus
florida, Pleurotus cornucopiae, and Pleurotus eryngii” in Mushroom Science IX (Part I). The
Mushroom Research Institute, Japan.

F. Zadrazil, 1978. “Cultivation of Pleurotus” in The Biology and Cultivation of Edible Mush-
rooms ed. by S.T. Chang and W.A. Hayes. Academic Press, New York.

I. Heltay, 1980. “Pleurotus florida Production in Borota, Hungary”. Mushroom Journal,
London.

SPECIES: Psilocybe cubensis (Earle) Singer
= Stropharia cubensis Earle.

= Stropharia cyanescens Murr.

= Stroparia caerulescens (Pat.) Sing.

= Naematoloma caerulescens Pat.

= Hypholoma caerulescens (Pat.) Sacc. & Trott.

Figure 163 Psilocybe cubensis fruiting on cased grain.

STRAINS: Strains of Psilocybe cubensis are available from private and commercial stocks. The
American Type Culture Collection, which sells cultures to educational organizations and research
facilities, has stock cultures of several wild strains. Note that the strains listed below are only some of
those that are presently circulating. There are many more. Some strains may originate from the
same region but have features not in agreement with those described here.

Amazonian: Medium to large mushrooms on rye grain; thick whitish stems; tenaciously at-
tached to the casing.

Growing Parameters for Various Mushroom Species/ 1 97

Ecuadorian: Medium sized mushrooms on rye grain; hemispheric caps; abundant primordia
former; high yielding on compost; thin whitish stems; easily picked.

Mafias Romero: Medium to large mushrooms on rye grain; early fruiter; thick whitish stems
and tenaciously attached.

Misantla: Medium sized mushrooms on rye grain; thin yellowish stems; tall standing and easi-
ly picked.

Palenque: Large mushrooms on rye grain; high yielding; and easily picked.

COMMON NAMES: San Isidro; Cubensis.

GREEK ROOT: Psilocybe comes from the Greek root “psilos” meaning bald head and cubensis,
a name Earle assigned to this mushroom because it was first recognized as a new species from
specimens collected in Cuba.

GENERAL DESCRIPTION: A medium to large size mushroom having a cap that becomes con-
vex to plane in age and is usually pigmented chestnut brown to deep yellowish or golden brown.
The cap surface is finely fibrillose, sometimes covered with scattered, fugacious, cottony scales that
soon disappear. The partial veil is membranous, well developed and typically leaving a persistent
annulus on the upper regions of the stem. The stem is often longitudinally striate, powdered above
the annulus and often covered with dense fibrils below. Flesh bruising bluish or bluish green. Its
spores purplish brown in mass.

NATURAL HABITAT: Naturally found in horse and cow pastures, in dung or in soil enriched with
manure. Psilocybe cubensis is a widely distributed species that is found throughout tropical and
subtropical zones of the world and is common in the pasturelands of the gulf coast of the southern
United States and eastern Mexico.

GROWTH PARAMETERS

Mycelial Types: Rhizomorphic to linear; whitish in overall color but often bruising bluish where in-
jured.

Standard Spawn Medium: Rye grain. See Chapter ill.

Fruiting Substrate: Rye grain; wheat straw; leached horse or cow manure; and/or horse
manure/straw compost balanced to a 71-74% moisture content.

Method of Preparation: See Chapters III, V, and VI. Pasteurization achieved through exposure to
live steam for 2 hours at 140°F. throughout the substrate. Straw or compost should be filled to a
depth of 6-12 inches. Straw should be spawned at a rate of 2 cups/sq. ft.

Spawn Run:

Relative Humidity: 90%.

Substrate Temperature: 84-86 °F. Thermal death limits have been established at 106°F.
Duration: 10-14 days.

C0 2 : 5000-10,000 ppm.

198/The Mushroom Cultivator

Fresh Air Exchanges: 0 per hour.

Type of Casing: After fully run, cover with the standard casing whose preparation is described in
Chapter VIII. Layer to a depth of 1-2 inches. The casing should be balanced to an initial pH of
6. 8-7. 2.

Post Casing/Prepinning:

Relative Humidify: 90 + %.

Substrate Temperature: 84-86 °F-
Duration of Case Run: 5-10 days.

C0 2 : 5000-10,000 ppm.

Fresh Air Exchanges: 0 per hour.

Light: Incubation in total darkness.

Primordia Formation:

Relative Humidify: 95-100%.
Air Temperature: 74-78 °F.

Figure 164 Psilocybe cubensis fruiting on cased straw.

Growing Parameters for Various Mushroom Species/ 1 99

Duration: 6-10 days.

C0 2 : less than 5000 ppm.

Fresh Air Exchanges: 1 -3 per hour.

Light: Diffuse natural or exposure for 1 2-1 6 hours/ day of grow-lux type fluorescent light high
in blue spectra at the 480 nanometer wavelength. (See Chapters IV and IX).

Cropping:

Relative Humidity: 85-92%.

Air Temperature: 74-78 °F.

C0 2 : less than 5000 ppm.

Fresh Air Exchanges: 1 -3 per hour.

Flushing Pattern: Every 5-8 days.

Harvest Stage: When the cap becomes convex and soon after the partial veil ruptures.

Light: Indirect natural or same as above.

Yield Potential: Average yields are 2-4 Ibs./sq.ft. over a 5 week crbpping period. Maximum yield
potential has not been established.

Moisture Content of Mushrooms: 92% water; 8% dry matter.

Nutritional Content: Not yet established.

Comments: One of the easiest mushrooms to grow, this species fruits on a wide variety of sub-
strates within broad environmental parameters. As a primary and secondary decomposer, Psilocybe
cubensis fruits well on untreated pasteurized straw and on horse manure/straw composts trans-
formed by microbial activity. Sterilized grain typically produces smaller mushrooms than bulk sub-
strates. Given the numerous substrates that support fruitings, Psilocybe cubensis is well suited for
home cultivation.

Psilocybe cubensis cultivation was unheard of twenty years ago. Today, this species ranks
amongst one of the most commonly cultivated mushrooms in the U.S. and soon the world. This
sudden escalation in interest is largely due to the publication of several popular guides illustrating
techniques for its culture.

Psilocybe cubensis is a mushroom with psychoactive properties, containing up to 1 % psilocybin
and/or psilocin per dried gram. The function of these serotonin-like compounds in the life cycle of
the mushroom is not known.

Genetic Characteristics: Basidia tetrapolar (4-spored), forming haploid spores (1 N); heterothal lie.
The mating of compatible monokaryons often results in fruiting strains. Clamp connections are
present. See Chapter XV.

For further information consult:

Oss, O.T. and O.N. Oeric, 1 976. “Psilocybin: Magic Mushroom Grower’s Guide’’. And/Or
Press, Berkeley.

200/The Mushroom Cultivator

SPECIES: Psilocybe cyanescens Wakefield

= Geophila cyanescens (Maire) Kuhn. & Romagn.
= P si locy be mairei Singer

Figure 165 Psilocybe cyanescens fruiting indoors in a tray of alder chips.

STRAINS: St. Clair.

Many wild strains can be adapted to cultivation.

COMMON NAMES: Cyan; Grandote.

GREEK AND LATIN ROOTS: Psilocybe comes from the Greek “psilos” or bald head. The
species name cyanescens is from “cyaneus” or blue for the color reaction of the flesh upon bruis-
ing.

GENERAL DESCRIPTION: Cap 20-50 mm, broad, convex to broadly convex to plane in age
with an elevated and undulating margin which is, in turn, translucent-striate. The cap surface is
smooth and viscid when moist from a separable gelatinous pellicle (“skin”). The color is caramel

Growing Parameters for Various Mushroom Species/201

brown, fading to yellow-brown to straw colored from the center. The gills are attached in an adnate
to adnexed fashion, dull brown with whitish edges. The stem is 60-80 mm. long by 2-5 mm. thick,
fibrous and enlarged towards the base. Its surface is smooth or powdered (pruinose). The stem color
is whitish, silky and becomes blue where injured, with rhizomorphs protruding about the stem base.
The partial veil is cortinate (cobweb-like), leaving little or no trace on the stem. Its spore print is dark
purplish brown.

NATURAL HABITAT: Clustered in woody habitats; in soils high in the tissue of deciduous trees;
or in tall rank grass. This species grows throughout the Pacific Northwest in areas well mulched by
woody debris of deciduous and coniferous trees (typically not associated with bark). It has been
reported from England and is thought to be broadly distributed throughout the European continent.

GROWTH PARAMETERS

Mycelial Types: Rhizomorphic to closely linear; whitish in color.

Spawn Medium: Sawdust/bran or rye grain spawn.

Fruiting Substrate: A lignicolous species utilizing a number of wood types, most notably alder,
maple and fir. It is able to grow on a wide variety of cellulosic wastes including newspaper and card-
board.

Method of Preparation: Branches and other small diameter wood are chipped into 1-3 inch
pieces, preferably in the spring when the sap content is highest. This material is spawned with
sawdust/bran (4:1) and made into prepared beds outdoors amongst ornamental shade plants
(especially rhododendrons) or tall grass. Another method is to use sawdust/bran or rye grain spawn
to inoculate soaked corrugated cardboard. When fully colonized, sheets of cardboard are laid at the
bottom of trays which are then covered with a 2-4 inch layer of freshly cut alder chips. (Wood chips
are far superior to sawdust as a fruiting substrate).

Spawn Run:

Substrate Temperature: 65-75 °F.

Duration: 30-60 days.

Relative Humidity: 90 + %

C0 2 : 10,000 ppm or higher.

Fresh Air Exchanges: 0 per hour.

Type of Casing: None required.

Primordia Formation

Relative Humidity: 95%.

Air Temperature: 50-60 °F.

C0 2 : 5000 ppm or below.

Fresh Air Exchanges: 2 per hour.

Light requirements: Diffuse natural or grow-lights.

202/The Mushroom Cultivator

Figure 166 Psilocybe cyanescens mycelium growing on soaked corrugated cardboard
inoculated with grain spawn.

Cropping:

Relative Humidity: 85-92%.

Air Temperature: 50-60 °F.

C0 2 : 5000 ppm or below.

Fresh Air Exchanges: 2 per hour.

Harvest Stage: When the caps become nearly plane.

Light: Diffuse natural or grow-lights.

Yield Potential: In natural outdoor culture on alder chips, 1 lb. wet weight per square foot in one
growing season is easily obtained.

Moisture Content: 90-92% water; 8% dry matter in fruitbodies.

Comments: Psilocybe cyanescens in a primary decomposer, readily digesting newly cut alder and
other deciduous woods. Considered the grandote of the Pacific Northwest, this species is both

Growing Parameters for Various Mushroom Species/203

robust and potently psilocybian. Much sought after for its high psilocybin and psilocin content, it is a
favored mushroom by those seeking entheogenic experiences.

Psilocybe cyanescens’ adaptability to natural outdoor culture makes this species attractive to
beginning and connoisseur cultivators alike. Virgin spawn can be collected from the wild and im-
planted in prepared beds (see Chapter VI) or spawn can be grown out on bran/sawdust or grain
and inoculated directly onto unsterilized soaked corrugated cardboard. Grain spawn inoculated onto
untreated wood chips is associated with a higher contamination rate than the same spawn implanted
onto soaked cardboard, owing to the partial selectivity of the latter material.

Although fruitbodies can form on fresh sawdust, they do so reluctantly and belatedly. The fact
that sawdust so readily loses its moisture may explain, in part, why Psilocybe cyanescens has diffi-
culty fruiting on it.

Psilocybe cyanescens has a mycelium that is typically whitish and strandy (rhizomorphic). Tis-
sue and spore cultures are easy to obtain. Outdoor colonies can be maintained for years with mini-
mal effort and produce two to three flushes within a season.

See Color Photos 17 & 18.

Figure 167 Sclerotia of Psilocybe mexicana harvested from one cup of rye grass seed
six weeks after inoculation.

STRAINS: Heim Strain
Pollock Strain

COMMON NAMES: Mushroom of the Gods; Teonanacatl or God’s Flesh; Nize (Mazatec Name);
and Pajaritos (Spanish Name).

GREEK ROOT: Psilocybe comes from the Greek “psilos” or bald head. The species name
mexicana denotes the country in which this mushroom grows.

GENERAL DESCRIPTION: Convex to subumbonate, sometimes with a small umbo, expanding
in age to plane or nearly so. The surface is smooth, translucent-striate two thirds to the disc. The cap
color is brownish to orangish grey to straw brown, more yellowish to the disc. The gills are adnately
attached, grey to dark purplish brown. The stem is equal, smooth, hollow, pale straw to brown to
reddish yellow, darkening when injured but typically not bruising bluish. Its spores are dark violet
brown in mass.

NATURAL HABITAT: Solitary to numerous in grassy areas, horse pastures and meadows al-

Growing Parameters for Various Mushroom Species/205

Figure 168 Two quart jars at 10 days and 30
days after inoculation onto rye grass seed.

though not occurring on dung. Distributed throughout subtropical regions in Mexico, common in
the state of Oaxaca, and also known from Guatemala.

GROWTH PARAMETERS

Mycelial Types: Slightly rhizomorphic to finely linear; off-white to tan in color, sometimes with
multicolored zones.

Spawn Media: Annual rye grass seed or rye grain.

Fruiting Substrates: Rye grass seed and to a lesser degree rye grain and pasteurized wheat straw.
Few fruitbodies form on enriched malt agar media.

Method of Preparation: Rye grass seed combined with water in a 2:1 volumetric proportion,
preferably soaked overnight and then sterilized for 1 hour at 1 5 psi. Wheat straw is pasteurized in a
hot water bath at 1 60-1 70 °F. for 30 minutes.

Spawn Run:

Relative Humidity: 90 + %

Substrate Temperature: 75-81 °F.

Duration: 10-1 4 days.

Relative Humidity: 90 + %.

COy. 10,000 ppm or higher.

Fresh Air Exchanges: 0 per hour.

Type of Casing: Standard peat based casing whose preparation is described in Chapter VIII. Layer
to a depth of 1 / 2-1 inch.

206/The Mushroom Cultivator

Post Casing/Pre-pinning:

Relative Humidity: 90 + %.

Substrate Temperature: 75-81 °F.

C0 2 : 10,000 ppm or above.

Fresh Air Exchanges: 0 per hour.

Light: Incubation in darkness.

Primordia Formation:

Relative Humidity: 95 + %.

Air Temperature: 71-74°F.

C0 2 : 5,000 ppm or below.

Fresh Air Exchanges: 2 per hour.

Light: Diffuse natural or fluorescent grow-lights for 12 hours daily.

Cropping:

Relative Humidity: 85-92%.

Air Temperature: 71-74°F.

C0 2 ‘. 5000 ppm or below.

Fresh Air Exchanges: 2 per hour.

Harvest Stage: When the caps become nearly plane.

Light: Same as above.

Yield Potential: Not yet established. A petite mushroom. Psilocybe mexicana is an interesting
species for the connoisseur. Because of the small stature of the fruitbody, one should expect low
yields per square foot. Sclerotia formation on rye grass seed after two months is 50-70 grams per
cup of seed.

Moisture Content: 90-92% water and 8% dry matter in fruitbodies; 70% water and 30% dry
matter in sclerotia.

Comments: This species is most remarkable for its early formation of sclerotia — only three weeks
after inoculation onto rye grass seed. Heim and Wasson (1 958) considered sclerotia production in
this species to be the most efficient method for the generation of biomass. Optimum temperature for
sclerotia production was reported to be at 70-75 °F. in darkness. Sclerotia on agar media peaked at
4.5% malt concentration. Heim and Wasson also found fruitbody production was maximized on
agar media when the percentage of malt was balanced to .45%. Nevertheless, sclerotia form best
on rye grass seed incubated in total darkness.

For further information consult:

“Les Champignons Hallucinogenes du Mexique” by R. Heim and R. G. Wasson, 1 958. Edi-
tions du Museum National D’Histoire Naturelle, Paris.

See Color Photographs 2 and 13.

Growing Parameters for Various Mushroom Species/207

SPECIES: Psilocybe tampanensis Guzman and Pollock

Figure 169 Sclerotia of Psilocybe tampanensis harvested from
one cup of rye grass seed six months after inoculation.

208/The Mushroom Cultivator

STRAINS: Pollock Strain.

COMMON NAMES: The Tampa Psilocybe; Pollock’s Psilocybe. Sclerotia are called The New
Age Philosopher’s Stone or Cosmic Comote.

GREEK ROOT: Psilocybe comes from the Greek “psilos” or bald head. The species name
tampenensis denotes the city near which this mushroom was first collected.

GENERAL DESCRIPTION: Cap convex to subumbonate, soon broadly convex to plane. The
surface is smooth and the color is ochraceous brown to straw brown to grey brown. The gills are ad-
nately attached, dark violet brown with whitish edges. The stem is 20-60 mm. long by 3-5 mm.
thick, fibrous and enlarged towards the base. The stem surface is smooth to powdered (pruinose)
and its color is yellowish brown to reddish brown overall, with whitish to bluish mycelium at or
around the base. Its spores are dark purplish brown in mass.

NATURAL HABITAT: Solitary to scattered in sandy soils and meadows in Florida (near the city of
Tampa). This species is known only from the type locality where one wild specimen was collected.

GROWTH PARAMETERS

Mycelial Types: Finely linear to cottony; tan to brownish in color, often multicolored with brownish
hues.

Spawn Media: Annual rye grass seed, wheat grass seed or rye grain.

Fruiting Substrate: Cased rye grass seed (and possibly rye grain); leached cow manure; some
potting soils; and enriched malt agar media. This species will probably fruit on cased pasteurized
wheat straw.

Method of Preparation: Rye grass seed combined with water in a 2:1 volumetric proportion,
preferably soaked overnight. Sterilize for 1 hour at 1 5 psi. Wheat straw is pasteurized in a hot water
bath at 160-1 70 °F. for 20 minutes.

Spawn Run:

Relative Humidity: 90 + %.

Substrate temperature: 75-81 °F.

Duration: 10-14 days.

C0 2 : 10,000 ppm or higher.

Fresh Air Exchanges: 0 per hour.

Type of Casing: Standard peat based casing whose preparation is described in Chapter VIII. Layer
to a depth of Zi to 1 inch.

Post Casing/Pre-pinning:

Substrate temperature: 75-81 °F.

Relative humidity: 90 + %.

C0 2 : 10,000 ppm or above.

Fresh Air Exchanges: 0 per hour.

Growing Parameters for Various Mushroom Species/209

Light: Incubation in darkness.

Primordia Formation:

Relative Humidify: 85-92%.

Air Temperature: 71-74°F.

C0 2 : 5000 ppm or below.

Fresh Air Exchanges: 2 per hour.

Light requirements: Diffuse natural or grow-iights for 1 2 hours/ day.

Cropping:

Relative humidity: 85-92%.

Air Temperature: 71-74°F.

C0 2 : 5000 ppm or below.

Fresh Air Exchanges: 2 per hour.

Harvest Stage: When the caps become nearly plane.

Light requirements: Diffuse natural or grow- lights for 12 hours/day.

Yield Potential: A petite species, Psilocybe tampanensis is noted for its sclerotia forming ability,
approximately 1 0-30 grams (wet weight) per cup of rye grass seed over 1 2 weeks. Because of the
small stature of the fruitbody, one should expect low yields per square foot in comparison to other
more fleshy species of Psilocybe.

Moisture Content: 90-92% water and 8- 1 0% dry matter in fruitbodies; 70% water and 30% dry
matter in sclerotia.

Comments: This mushroom would not be known but for a single specimen collected by Steven
Pollock and Gary Lincoff in September of 1 977. Cultures taken from this wild specimen were mar-
keted by Hidden Creek Inc. under the name of the “Cosmic Comote”.

Sclerotia do not form until the fourth week (typically six to eight weeks) after inoculation of rye
grass seed. To encourage sclerotia production only, incubate mycelia on rye grass seed at 75 °F. in
complete darkness.

For further information consult:

“Magic Mushroom Cultivation” by Steven H. Pollock, 1977. Herbal Medicine Research
Foundation, San Antonio, Texas (out of print).

See Color Photographs 4, 14 and 15.

21 0/The Mushroom Cultivator

SPECIES: Stropharia rugoso-annulata Farlow apud Murriit
= Stropharia ferii Bresadola
= Naematoloma ferii (Bres.) Singer

> ■ •* .

Figure 170 oung fruitbodies of Stropharia rugoso-annulata fruiting on pasteurized
straw cased with peat.

STRAINS: Gartenriese
Winnetou
Gelbschopf

The above listed strains are of European origin. Many strains of this species are available from
culture banks, including those maintained by the American Type Culture Collection and
Pennsylvania State Buckhout Laboratory. Strains are easy to obtain from the spores and tissue of
wild specimens.

COMMON NAMES: The Wine Red Stropharia; The Giant Stropharia.

LATIN ROOT: Stropharia means “sword belt”, so named for the belt-like ring on the stem. The
species epithet rugoso-annulata comes from the combination of two Latin words: “rugosus mean-
ing wrinkled and “annulus” or ring.

Growing Parameters for Various Mushroom Species/21 1

Figure 171 Buttons of Stropharia rugoso-annulata
fruiting outdoors in a bed of wood chips.

GENERAL DESCRIPTION: A large, thick fleshed mushroom with a broadly convex cap measur-
ing 50-400 mm. in diameter, darkly pigmented yellowish brown with distinct reddish tones. The
partial veil is thick, membranous, leaving a persistent membranous ring on the stem on whose up-
persides are tiers of gills. The stem is whitish and has rhizomorphs attached to its base. Its spores are
dark purplish brown in mass.

NATURAL HABITAT: Occurring in gardens, in wood chips, on decomposing straw, in sawdust
enriched soils and commonly in grounds where potatoes have been planted.

GROWTH PARAMETERS

Mycelial Types: Rhizomorphic to closely linear; whitish in color.

Standard Spawn Media: Rye grain or chopped wheat straw.

Fruiting Substrates: Cased wheat straw, whole or chopped, and balanced to a 71 -74% moisture
content. This species has been grown on a substrate of alder/ maple chips mixed with mature horse
manure using natural culture techniques.

212/The Mushroom Cultivator

Method of Preparation: Either chopped or whole straw is adequate, although permeation is more
rapid on the former. (See Chapter VI on preparation of straw as a fruiting substrate). Pasteurization
is achieved through the submersion of straw into a hot water bath at a temperature of 160°F. for
20-30 minutes. The straw, once pasteurized and inoculated, should be compacted and filled to a
depth of 6-1 2 inches. Gramss (1 979) noted that wheat straw supplemented with 25% Fagus saw-
dust enhanced yields. Watling (1 980) reported, without elaboration, that fruitbodies form on a saw-
dust based medium. This species also fruits on unpasteurized straw although problems with insect
pests and competitor molds are more pronounced.

Spawn Run:

Relative Humidify: 90 + %.

Substrate Temperature: 76-82 °F. Thermal death limits have been reported as low as 90 °F.
and — 5°F.

Duration: 2-4 weeks.

C0 2 : 5000-10,000 ppm.

Fresh Air Exchanges: 0 per hour.

Type of Casing: After fully run, cover with peat/humus (1:1) casing. Optimally, the casing should
have a pH of 5. 7-6.0. (Because calcium based buffers inhibit fruiting, adjust the casing’s pH by in-
creasing or decreasing amount of peat). Balance to a 70-75% moisture content. Layer to a depth of
1-2 inches. Humus should be pasteurized to kill nematodes, mites, and other parasites. Some
strains form fruitbodies solely on a peat casing. (Mushrooms do not form, however, on sterilized
casing. Hence, if the casing must be treated, steam pasteurization is recommended).

Post Casing/Prepinning:

Relative Humidify: 90 + %.

Bed Temperature: 76-82 °F.

Duration of Case Run: 10-12 days.

C0 2 : 5000-10,000 ppm.

Fresh Air Exchanges: 0 per hour.

Light: Incubation in darkness.

Primordia Formation:

Relative Humidify: 95 + %.

Air Temperature: 55-62 °F.

C0 2 . less than 1000 ppm.

Fresh Air Exchanges: 2-4 per hour.

Watering: Regular misting (once to twice daily) to help stimulate primordia formation.

Light: Indirect natural or exposure to grow-lux type fluorescent for 12 hours/day.

Cropping:

Relative Humidify: 85-92%.

Growing Parameters for Various Mushroom Species/21 3

Air Temperature: 55-62 °F.

C0 2 '■ less than 1000 ppm.

Fresh Air Exchanges: 2-4 per hour.

Flushing Interval: Every 10-15 days.

Harvest Stage: Directly before or as the partial veil tears. (Note that young mushrooms have a
much better flavor than mature ones).

Light: Indirect natural or exposure to grow-lux type fluorescent for 12 hours/day.

Yield Potential: Average commercial yields are 2-3 lbs./sq.ft. over a 8 week cropping period.
Maximum yields are nearly 6 lbs per square foot.

Moisture Content of Mushrooms: 92% water; 8% dry matter.

Nutritional Content: 22% protein (dry weight); 34 milligrams of niacin per 100 grams dry
weight.

Comments: A mushroom recently cultivated in Europe (Germany, Czechoslovakia and Poland)
by home growers in outdoor cold frames, the status of knowledge regarding the optimum growing
parameters for this species remains in its infancy. For instance, Szudyga (1978) noted that fruitbod-
ies form just as well at 50 °F. and 68 °F., a considerable fruiting range for any species.

After the cropping period ends, the spent straw is used as fodder for farm animals or is saved for
future inoculations. The strain is kept kept alive by continous transfer onto fresh substrates. (See
Chapter VI on natural culture). Propagating spawn in this way, however, is less assured than sterile
methods.

Stanek (1974) reported that the introduction of several thermotolerant endospore-forming bac-
teria of the genus Bacillus (B. subtilus, B. meseatericus and B. macerans) to the casing not only in-
hibited attacks by competitors but also stimulated mycelial growth which presumably would en-
hance yields. Endospores of these bacteria survive pasteurization but not sterilization, and are abun-
dant in soils. This discovery may explain why sterilized casings do not produce fruitbodies.
Genetic Characteristics: Basidia tetrapolar (4-spored), forming haploid spores; heterothallic.
Clamp connections are present. See Chapter XV.

For further information consult:

K. Szudyga, 1978. “Stropharia rugoso-annulata” in The Biology and Cultivation of Edible
Mushrooms ed. by S.T. Chang and W.A. Hayes. Academic Press, New York.

214/The Mushroom Cultivator

SPECIES: Volvariella volvacea (Bull, ex Fr.) Sing.

STRAINS: Many strains of V. volvacea are available from commercial and private stocks. The
American Type Culture Collection, which sells cultures to educational organizations and research
facilities, has stock cultures of several wild and domesticated strains. Several commercial companies
also sell strains of this species.

COMMON NAMES: The Paddy Straw Mushroom; The Chinese Mushroom.

LATIN ROOT: Volvariella is the conjunction of two words: “volvatus” which means having a
volva or cup-like sheath and the suffix “-ellus” denoting smallness in size. The species name
volvacea shares the same root as the genus.

GENERAL DESCRIPTION: Mushrooms whitish at first, becoming a dark tan as the veil tears and
eventually a pale tan with age. Fruitbodies are relatively small when young, enveloped by a sheath-
like universal veil, soon breaking as the fruitbodies mature and leaving an irregular cup-like sack at
the base of the stem. The cap is egg shaped at first, soon hemispherical to convex and expanding to
plane with age. Its spores are pinkish to pinkish brown in mass.

NATURAL HABITAT: Commonly occurring in decomposing straw in the Orient and in other
subtropical regions of the world.

GROWTH PARAMETERS

Mycelial Types: Fast growing rhizomorphic to slow cottony mycelia noted. The color is typically
white to grayish white.

Spawn Medium: Rice straw or rye grain. See Chapter III.

Fruiting Substrate and Method of Preparation: Traditionally grown on rice straw that has been
composted for 1-2 days. More recently Hu (1974) found that a mixture of cotton wastes supple-
mented with wheat bran and calcium carbonate (5% and 5-6% by weight, respectively) and com-
posted for 3 days, pasteurized for 2 hours at 1 40 ° F., conditioned for 8 hours at 1 25 0 F. and then
gradually lowered to 77 °F. over a 8-1 2 hour period, produced a higher yielding substrate than that
of others previously used. A moisture content of 65-70% is recommended for rice straw and 70%
for cotton waste mixtures. Chang (1978) recommended a combination of the two— with the rice
straw/ cotton waste in a proportion of 2:1 or 1:1 by weight.

Spawn Run:

Relative Humidity: 90 + %.

Substrate Temperature: Fastest growth at 88-95 °F.

Duration: 4-6 days for thorough colonization.

C0 2 : 5000-10,000 ppm.

Fresh Air Exchanges: 0 per hour.

Light Requirements: Incubation in total darkness.

Type of Casing: None needed.

Growing Parameters for Various Mushroom Species/215

Pinhead Initiation:

Relative Humidity: 95 + %.

Air Temperature: 82-88 °F.

Duration: 4 days.

C0 2 : less than 1000 ppm.

Fresh Air Exchanges: 2-4 per hour.

Light: Diffuse natural or direct grow-light fluorescent for 12-18 hours per day.

Watering: Regular misting once to twice daily.

Cropping:

Relative Humidity: 85-92%.

Air Temperature: 82-88 °F.

Duration: 5-7 weeks.

C0 2 : less than 600 ppm.

Fresh Air Exchanges: 2-4 per hour or sufficient to meet C0 2 and/ or cooling requirements.
Harvest Stage: Directly before rupturing of the universal veil.

Flushing Intervals: 5-10 days.

Light: Same as above.

Watering: Regular misting to prevent caps from cracking and to keep resting pinheads viable.
Yield Potential: Average commercial yields on rice straw are 22-28 kilograms of fresh mush-
rooms per 100 kilograms of dry straw. Optimum yields on cotton waste compost are 25-35
kilograms per 100 kilograms of substrate. Maximum yields are nearly 45 kilograms on cotton
waste compost.

Moisture Content of Mushrooms: 88-90% water; 10-12% dry matter.

Nutritional Content: Crude protein is reported at 21 .2 % of dry weight; 91 milligrams of niacin
per 1 00 grams dry weight.

Comments: In contrast to other species growing on straw, this mushroom does not compare fa-
vorably in terms of yield. The smaller crop figures are probably a result of the early picking of the
mushroom fruitbodies, when they are most flavorful.

Several researchers have noted the difficulty of maintaining high yielding strains of this species
for any length of time. Its mycelium seems to have a limited transfer potential and should be stored
at moderate temperatures (50 °F.). Cultures are frequently renewed through multispore germina-
tions.

Volvariella volvacea is primarily grown in the Orient and is a warmth loving mushroom.
Genetic Characteristics: Basidia tetrapolar, producing 4 haploid spores; primary homothallic.
Clamp connections are present. Chlamydospores form. See Chapter XV.

For more information consult:

S.T. Chang, 1972. “The Chinese Mushroom ( Volvariella volvacea): Morphology, Cytology,

216/The Mushroom Cultivator

Genetics, Nutrition and Cultivation'’ The Chinese University of Hong Kong, Hong Kong.

S.T. Chang, 1978. “Volvariella votvacea” in The Biology and Cultivation of Edible Mush-
rooms, pp. 573-603. Academic Press, New York.

Cultivation Problems and Their Solutions/217

Figure 172a,b,c,d. The results of bacterial contamination

■ sl

i> j

218/The Mushroom Cultivator

M any first-time cultivators fail to grow mushrooms for the simplest of reasons. Often times the
slightest error in technique sets into motion a series of events that drastically influence the out-
come of the crop. Whenever conducting sterile technique, making spawn, preparing compost or
cropping mushrooms, wise cultivators follow a routine that has proven successful in the past. Once
a consistent methodology has been established, new variations are introduced, one at a time, to
gauge their effect.

Problems intrinsic to mushroom culture have been encountered by most everyone attempting
to grow mushrooms. The following trouble-shooting guide lists problems, causes and solutions ac-
cording to their frequency of occurence and has been organized into five categories:

1 . Sterile Technique: media (agar and grain) preparation, spore germination, tissue culture
and spawn-making.

2. Compost Preparation: raw materials, characteristics of composts at differents stages,
Phase I and Phase II.

3. Spawn Running: colonization of compost and bulk substrates.

4. Case Running: application, colonization by mycelium, pre-pinning strategy.

5. Mushroom Formation and Development (Pinning to Cropping): strategy for pinhead
formation, maturation and harvesting.

Identify the problem, locate it on the list, read its possible causes, refer to the solutions availa-
ble, and if indicated, turn to the chapter noted in parentheses. Good luck, pay attention to detail and
may your problems be few.

Cultivation Problems and Their Solutions/219

STERILE TECHNIQUE

PROBLEM

CAUSE

SOLUTION

Agar Culture

Media fails to solidify.

Insufficient quantity of agar or
distribution thereof.

Thoroughly mix media before
pouring.

Media boils out of vessel or
flask containing it.

Excessive escape of steam
from pressure cooker.

Do not vent pressure cooker
until reaching 1 psi.

Grease pressure cooker seals
with thin film of petroleum jel-
ly.

For pressure cookers using 5,
10, 15 lb weights, do not
operate so steam escapes.

Contamination occurs in petri
dishes after pouring media
but before inoculation.

High contaminant spore load
in lab.

Clean, paint lab. Install lami-
nar flow hood.

Improper media preparation
technique.

Contaminated pressure
cooker (bacteria).

Allow pressure cooker to cool
in sterile setting before open-
ing.

Sterilize pressure cooker for
24-48 hours at 1 5 psi.

No growth from spores or tis-
sue transferred.

Wrong type of media.

Wrong pH.

Old or dehydrated spores.
Scalpel or loop too hot.
Sugar in media caramelized.

See media preparation.

See media preparation
Soak in sterilized water for
1 2-24 hours.

Cool tool before contacting
spores or tissue.

Lower sterilization pressure

and temp, to recommended
levels.

220/The Mushroom Cultivator

PROBLEM

CAUSE

SOLUTION

Contamination occurs atound
point of transfer onto agar
media.

Inoculum (spores or tissue)
contaminated.

Inoculation tools not sterile.

Obtain “cleaner” spores or
take a tissue culture from a
fresher specimen, or inoculate
as many plates as possible,
saving only those not becom-
ing contaminated.

Autoclave tools, soak in alco-
hol, flame sterilize before us-
ing.

Rhizomophic mycelia be-

Senescence, strain aging.

Retrieve stock cultures and re-

comes cottony, slow growing.

activate a strain of known

Fruitings diminish. Strain ap-

vigor.

pears to be degenerating.

Mutating. Sugar in media car-

Alternate media so that gene
expression is not selected by
a limited chemical matrix.

Cook agar media at lower

amelized. Media containing

temp, and pressure, between

mutagens.

1 2 and 1 5 psi.

Insufficient jelling agent caus-

Add more agar or thoroughly

ing mycelium to grow subsur-

mix media before pouring

facely and appear cottony.

petri dishes.

Grain Culture

Glass spawn jars broken
when pressure cooker is
opened.

Pressure cooker cooled too
rapidly. Change in temp, too
abrupt.

Jars too tightly packed.

Jars defective or cracked.
Wrong type of jars.

Allow cooker to descend to
room temp, gradually.

Allow space so jars can ex-
pand.

Check for cracks or defects.
Obtain new jars.

Replace with canning or auto-
clavable type.

Cultivation Problems and Their Solutions/221

PROBLEM

CAUSE

SOLUTION

Crain jars difficult to shake.

Too much grain in container.

Reduce grain to recom-
mended levels.

Too much water relative to

Follow recommended for-

grain.

mulas.

Measuring cups not accurate.

Calibrate measuring cups with
a graduated cylinder.

Grain “spontaneously” con-

Introduction of alien spores

Cool-down in sterile environ-

taminates before inoculating

upon cooling.

ment or in front of laminar

or opening pressure cooker.

flow hood.

Survival of bacterial endo-

Replace source of grain or

spores despite autoclaving.

presoak grain for 24 hours
before autoclaving.

Agar wedge sticks to glass

Agar media too thin, either

Use mycelium covered media

when grain jar is shaken.

from evaporation or from

before substantial evaporation

shallow pouring.

occurs. Pour more media into
each petri dish initially.

Little or no growth after my-

Crain too hot when inocu-

Allow to cool to room tern-

celial wedge has been trans-

lated.

perature before inoculating.

ferred.

Grain too dry.

Balance according to recom-
mended recipes.

Mycelium not evenly distrib-

Vigorously shake spawn jar

uted.

after transfer of agar wedge
and again 3-5 days after inoc-
ulation.

Incubated at wrong tempera-

See recommended spawn in-

ture.

cubation temperatures in
Chap. XI.

pH wrong.

Buffer with calcium carbonate
according to species being
cultured.

Wrong spawn medium.

Use media recommended for
that species.

222/The Mushroom Cultivator

PROBLEM

CAUSE

SOLUTION

Poor strain.

Contaminated strain.

Discard strain. Obtain purer
strain, make up more grain
media and clean laboratory.

No growth on grain after in-

Too many individual hyphae

Stirred for too long. No more

oculated with liquid cul-

(cells) severed.

than 5 seconds is recom-

ture/stirrer technique.

mend for high speed labora-
tory-type blenders to produce
fragmented chains of hyphae.

Bacteria.

Replace mycelia with pure
strain, free of bacteria. Be
sure tools and water are ster-
ile before inoculation.

Poor strain.

Cottony type mycelia is slow
growing. Replace with rhizo-
morphic or faster growing
strain.

Follow recommended recipes.
See Chap. II.

Water too hot.

Allow to cool before inoculat-

ing.

Contamination after transfer of

High contaminant spore

Clean lab before inoculations.

mycelium.

count in laboratory.

Maintain high standards of
hygiene.

Tools not sterile.

Autoclave tools, soak in alco-
hol, flame sterilize before in-
oculation.

Mycelium being transferred

Obtain cleaner strain or

has high resident load of con-
taminant spores.

spawn of better purity.

Mycelium fails to grow out

Insufficient shaking of grain

Thorough shaking after dou-

through entire spawn jar.

after inoculation.

ble-wedge transfer, combined
with re-shaking four days after

inoculation.

Cultivation Problems and Their Solutions/223

PROBLEM

CAUSE

SOLUTION

Mycelium inhibited by con-
tamination (usually bacteria).
Externally or internally intro-
duced.

Inoculate more sterilized grain
using a pure strain and fol-
lowing standard practices for
doing so. See Chap. II.

Top kernels in spawn jar not
colonized by mycelium.

Top kernels dehydrated from
excessive evaporation.

If using porous filter discs,
limit evaporation. Or use only
in conjunction with narrow
mouthed jars.

Spawn jar discolored with yel-
lowish droplets of fluid.

Spawn jar incubated for an
overly long period of time, at
higher than optimum temper-
atures, or both, causing the
exudation of metabolites
(“sweat”) and the build-up of
fluids in which bacteria thrive.

Incubate at temperature and
for period of time
recommended for species
being cultivated.

COMPOST PREPARATION

Phase f

Compost does not heat up,
remains under 140°F.

Undersupplemented.

Pile too open, airy.

Moisture content too high or
low.

Insufficient pile mass.

Check compost formula.
Check Nitrogen content of
raw materials.

Compress pile sides. Protect
pile from strong winds.

Balance moisture to 70%.
Increase total raw materials.

Compost generates no am-
monia.

Undersupplemented.

Check compost formula cal-
culations. Check Nitrogen
content of raw materials.

224/The Mushroom Cultivator

PROBLEM

CAUSE

SOLUTION

Compost anaerobic.

Moisture content too high,.
Straw too short; pile too
dense.

Pile sitting too long between
turns.

Balance moisture to 70%.
Carefully monitor raw materi-
als and adjust pile size as ma-
terials compact.

Turn more frequently.

Compost decomposing un-
evenly.

Improper turning procedures.
Variable starting materials.

Move inside of pile to outside
and vice versa.

Horse manure or straw
should ail be in the same
state of decomposition at the
start of composting.

Compost greasy.

Gypsum quantify too low.
Starting materials too old.

Add more gypsum.

Use only fresh, undecom-
posed starting materials.

Compost too wet or too dry
at filling.

Incorrect water addition or
timing.

Check moisture content of
pile before each turn.

Straws still bright and shiny at
filling.

Phase 1 too short.

Continue composting.

Compost short and black at
filling.

Phase 1 too long.

Shorten Phase 1.

Phase I!

Compost will not heat up.

Supplementation rates too
low.

Compost too mature.
Oversupply of fresh air.
Air to bed ratio too great.

Check compost formula cal-
culations.

Shorten Phase I.

Reduce fresh air supply.

Add more beds or trays and
fill with more compost.
Compost should be 70% at
filling.

Compost too wet.

Cultivation Problems and Their Solutions/225

PROBLEM

CAUSE

SOLUTION

Compost temperature erratic.

Irregular fresh air supply.

Room environment not moni-
tored enough.

Fresh air supply should be
constant. Make volume
changes slowly and as needed
to stablize temp.

Check room every 4-6 hours.

Compost temperature un-
even.

Containers filled unevenly.

Uneven supplement distribu-
tion in Phase 1.

Faulty air system design.

Fill all containers with equal
amounts of compost and to
the same depth.

Be sure supplements are
evenly mixed and are not
concentrated in small pockets.

Air system should insure even
temp, throughout the room.

Compost temp, too high after

Inadequate supply of fresh air.

Increase fresh air.

pasteurization.

Pasteurization too long.

Pasteurize for 2 hours at
1 40 °F.

Compost temp, drops too low
after pasteurization.

Prolonged fresh air supply.

Pasteurize at a lower temp,
for more time.

Anticipate drop in compost
temp, and reduce fresh air
before reaching conditioning
temp.

Low temperatures preserve
more microorganisms that
prevent temp, from falling
rapidly.

Prolonged ammonification.

Oversupplementation with
nitrogen.

Prolonqed time at temp, over
1 30 °F.

Reduce nitrogen supplements.

Keep temp, under 130° after
pasteurization.

Use low temp, ranges during
conditioning.

226/The Mushroom Cultivator

PROBLEM

SPAWN RUNNING

CAUSE

SOLUTION

Spawn grows slowly or not at
all.

Inferior spawn.

Degenerative or inviable
strain.

Check spawn making proce-
dures. Review strain storage
methods. Test strain purity by
inoculating agar plates.
Always test untried strains in
“miniculture” trials prior to
inoculation into bulk
substrates. Switch to a strain
of known viability.

Residual ammonia in com-
post.

Improper Phase 1 or Phase II.
Substrate moisture content
too high.

Fly or nematode infestation.
Mycelium lacks oxygen.

Prolong Phase II conditioning
until litmus paper test shows
no color change.

Review composting section.

Compost should be 64-66%
water; straw should be
70-75% at spawning.

Check pasteurization time and
temperature.

Be sure the container has
provisions for air exchange.

Molds present during spawn
run.

Improper Phase I or Phase II.

Review composting section.
Check contamination section
for identification and factors
predisposing to mold growth.

Inky Caps (Coprinus sp.) oc-
cur during spawn run

Residual ammonia in com-
post.

Prolong Phase II conditioning
until litmus paper test shows
no change.

Mites or nematodes present.

Insufficient pasteurization.
Compost with dense overwet
areas.

Unclean substrate containers
or spawning tools.

Pasteurize 2 hrs. at 140° F.
Review composting and filling
procedures.

Containers and tools should
be disinfected before use.

Cultivation Problems and Their Solutions/227

CASE RUNNING

CAUSE SOLUTION

PROBLEM

Mycelium fails to run through Temperature too high or too
the casing layer. low.

pH improperly adjusted.

Casing dries out after applica-
tion.

Casing to wet or too dry.

Unsatisfactory casing materi-
als.

Weak mycelial growth in sub-
strate.

Substrate contaminated.

Growing room humidity too
low.

Uneven mycelial growth into
substrate underlying casing.

Incubate at optimum temp,
for mycelial growth.

Test pH before application.
Adjust with limestone buffer
(with one exception). Consult
Chap. XI on the correct pH
for each species.

Test moisture before applica-
tion. Apply at 90% of capaci-
ty (70-75% moisture).

Review preparation of casing
in Chap. VIII.

Review techniques for sub-
strate preparation in appropri-
ate chapter.

Check substrate for molds
and nematodes before casing.

Increase humidification. In-
crease frequency of watering,
or cover with plastic.

Decrease fan speed. Maintain
slow, easy circulation.

Thoroughly mix casing ingre-
dients to insure an even
blend.

Redistribute or apply casing to
an even depth.

Thoroughly and evenly
spread spawn throughout sub-
strate at inoculation.

Fan speed too high. Too
much airflow.

t*.?

Uneven mycelial growth into
casing.

Casing mater|ajgjac|Mly
mixed. s?

\S G P

Unevenly applied casing.

228/The Mushroom Cultivator

PROBLEM

CAUSE

SOLUTION

Mycelium covers the casing
but forms few primordia.

“Overlay” caused by pro-
longed mycelial growth into
the casing layer.

Patch the casing. Begin initia-
tion sequence sooner. If deal-
ing with a slow pinning strain
be careful that the evaporation
rate off the casing surface is
not excessive.

Mycelium overlays the casing
and then “mats”, becoming
flattened and impervious to
water. No primordia form.

Improper watering and/ or too
low humidity in the external
environment. Evaporation rate
too extreme.

Scratch and/or re-case. Main-
tain 95% humidity at pinning.
Reduce evaporation rate. If
watering, mist lightly and
evenly.

Mycelium runs through the
casing and then disappears.

Die Back Disease (Virus).

Discard and begin anew with
a virus-free strain. See Chap.
XIII.

Dense white matted zones Stroma,
form on casing.

Contaminant (Scopulariopsis).

Select strains not predisposed
to stroma formation (those
without fluffy sectors). Reduce
C0 2 .

pH too high. Compost im-
properly prepared. See Chap-
ters V, XIII.

Cultivation Problems and Their Solutions/229

MUSHROOM FORMATION AND DEVELOPMENT

PROBLEM CAUSE SOLUTION

Pinhead Initiation

Mycelium fails to form pri-

Monokaryotic strain with low

Start again with new tissue

mordia.

or no fruiting ability.

isolate or isolate from multi-
spore germination.

Humidity too low.

Keep humidity at 95% during
pinning.

C0 2 too high.

Reduce C0 2 by introducing
fresh air.

Temperature too high.

Decrease air temp, to the
fruiting range.

Insufficient light.

Illuminate cropping surface
for 12 hours/day.

Primordia form early.

Uneven casing depth.

Apply casing at an even
depth and patch areas where
mycelium appears premature-
ly.

Incubate culture in darkness
until ready to pin.

Early light stimulation.

Temperature too low.

Incubate at optimum temp,
for mycelial growth and then
drop temp, for pinning.

C0 2 levels too low.

Maintain airtight room and re-
circulate air until ready to pin.

Primordia formation uneven.

Uneven casing depth.

Patch shallow areas as myce-
lium appears until growth is
even.

Uneven moisture in casing.

Water casinq evenly and care-
fully.

Casing surface partially dam-

Keep casing surface open and

aged from heavy watering.

porous through proper mist-
ing techniques.

Uneven environmental condi-
tions within the growing
room.

Review air system design.

230/The Mushroom Cultivator

PROBLEM

CAUSE

SOLUTION

Pinheads fail to form abun-

Casing layer moisture too low

Adjust moisture level in

dantly.

or too high.

casing to /U-/5Vo tor

pinhead formation.

Casing layer pH imbalanced.

Adjust pH to levels recom-
mended in Chap. XI for the
species being cultivated.

Magnesium in limestone buf-

Some species are inhibited by

fer too high (above 2%).

minerals in the casing layer,
especially the magnesium in
dolomitic limestone. Use a
low magnesium lime, less
than 2%.

C0 2 too high.

Lower C0 2 to recommended
levels. (Some species fruit
poorly in high C0 2 environ-
ments).

Insufficient light.

Photosensitive species require
several hours of light stimula-
tion per day for pinhead for-
mation.

Improper pinhead initiation
strategy.

See Chapter IX.

Defective strain.

Replace with strain of known
viability.

Nematode infestation.

See Chapter XIV.

Pinheads form but fail to ma-

Insufficient nutrient base.

Review substrate materials

ture.

and formulas. Follow those

that are recommended for the
species being cultivated.

Excessive C0 2 levels.

Reduce C0 2 to recom-
mended levels.

Humidity to high.

Reduce humidity to 85-92%.

Insufficient fresh air.

Increase fresh air input to 2-4
room exchanges per hour.

Strain idiosyncrasy.

Replace with a strain having
better fruiting calabilities.

Cultivation Problems and Their Solutions/ 231

PROBLEM

CAUSE

SOLUTION

Fly, nematode or other con-
taminant inhibiting develop-
ment.

Excessive loss of moisture
from casing.

Review contaminant control
procedures. Check source
and quality of casing materi-
als. Check mixing proce-
dures.

Maintain sufficient moisture
(70-75%) in the casing
through daily mistings if re-
quired.

CROPPING

Low yielding first flush.

Poor pin set.

Substrate low in nutrients.

Review pinning procedures
and growing parameters for
the species being grown.
Review substrate materials
and formulas.

Few mushrooms develop ful-
ly, many abort.

Uneven pinning.

Lack of nutrients.
Temperature too high.
Parasitized by contaminant.

Remove early developing
pins.

Review substrate materials
and formulas.

Maintain air temp, within
cropping range.

See Chapters X and XIII. Fol-
low procedures for encourag-
ing cropping, not contamina-
tion.

Mushrooms have long stems

C0 2 too high.

Increase fresh air input.

and small underdeveloped
caps.

Insufficient lighting.

Evaluate lighting system and
type of light used.

232/The Mushroom Cultivator

PROBLEM

CAUSE

SOLUTION

Mushrooms develop but ab-
normally.

Parasitized by contaminant.

Eliminate stagnant air pockets
in the growing environment.
See Chap. IV.

Excessive C0 2 . Improperly
balanced growing environ-
ment

Lower C0 2 to recommended
levels. Maintain air circulation,
temp, and humidity at recom-
mended levels. See Chap. X.

Exposure to mutagenic chem-
icals (insecticides, detergents,
chlorine, etc.)

Limit exposure of mushrooms
to such chemicals.

Lack of adequate light for
fruitbody development.

Increase light exposure to 12
hours per day. See Chapters
IV and IX.

Strain idiosyncrasy.

Switch to strain of known
fruiting ability.

The Contaminants of Mushroom Culture/233

3xr< ><-? jajg^r J

234/The Mushroom Cultivator

T he contaminants are so named solely because they are undesired. If one were trying to culture
Penicillium and spores of an Agaricus or Psilocybe settled onto the agar media and germi-
nated, the resulting mycelia would be the so-called “contaminant.” The contaminants in mushroom
culture, however, are primarily molds, bacteria, viruses and insects. The pathway by which a disease
is introduced, known as the vector of contamination, can be used to trace the contaminant back to
its site of origin using simple deduction. By observing how a contaminant affects the mushroom
crop and by carefully noting the conditions in which it flourishes, a cultivator can soon identify its
cause.

Earlier in the book, the five most probable vectors of contamination were identified as:

1. the cultivator.

2. the air.

3. the substrate to be inoculated.

4. the mycelium that was being transferred.

5. the inoculating tools, equipment, containers, facilities, etc.

Different contaminants are associated with different stages of mushroom cultivation. Contami-
nants in agar culture most often come from airborne spores. Grain cultures contaminate from air-
borne spores and from a source which many cultivators fail to identify: the grain used in spawn mak-
ing which is laden with spores of imperfect fungi, yeasts and bacteria. (See Ivanovich-Biserka,
1972). In compost culture, the major contributors to contamination are the materials used, the
spawn, the workers or the facilities. This is not to say that contaminants can not be introduced by
other means; these are the most probable sources of contamination given the cultivator has followed
generally accepted procedures for mushroom culture.

Tracking down the source of contamination is not difficult. For instance, the photographs be-
low show two media filled petri dishes contaminated with a Penicillium mold. Although the contam-
inant may be the same, the source of contamination is likely to be quite different. The plate in Fig.
174 has a mold colony growing directly beside the wedge of mycelium that was transferred. The
plate in Fig. 175 shows contamination along the outer periphery. Flere is a clear example illustrat-
ing how contamination spreads.

The left plate became contaminated when the mycelium was transferred, suggesting the mold
was associated with the previous culture. The right petri dish contaminated from airborne spores
which entered as the culture was incubating, judging by the proximity of the mold colonies to the
outer edge. Air movement within the “sterile” laboratory most likely wafted spores towards the
media plate and some penetrated the minute spaces belween the lid and the base. Within the still air
environment of the petri dish, spores settled nearest to their point of entry, germinated and began
resporulating, soon to be visible as a green mold. One would, therefore, implement the measures of
control accordingly.

Often times the source of contamination is not obvious. Beginners are at a particular disadvan-
tage because every contaminant they encounter is “new”. With each crop, problems arise requiring
novel solutions. If a certain method of cultivation has been repeatedly successful in the past and sud-

Figure 174 Penicillium mold near to Figure 175 Penicillium mold along outer

transferred wedge of mushroom myceli- periphery of petri dish.

um.

denly an unfamiliar contaminant appears, identifying the vector can be much more difficult. Only
when the cultivator can pinpoint the variables leading to the introduction of that contaminant can ap-
propriate counter-measures be applied. Frequently what seems to be an inconsequential alteration
in technique at one stage leads to a radical escalation of the contamination rate at later stages.

Since contamination at any phase of cultivation occurs for specific reasons, the contaminants
can be the cultivator’s most valuable guide for teaching one what NOT to do. If the problem causing
organism is identified and if the recommended measures of control are carefully followed, a con-
scientious cultivator will avoid those conditions predisposing to that one competitor and, incidental-
ly, many others. In effect, skill in mushroom culture is tantamount to skill in contamination control.

Molds and bacteria do not grow well in a climate specifically adjusted for mushrooms. Although
both mushrooms and contaminants prefer humid conditions, the latter thrive in prolonged stagnant
air environments whereas mushrooms do not. The differences are frequently subtle— amounting to
only a few percentage points in relative humidity and slight adjustments to the air intake dampers in
the growing room.

The contaminants can be divided into two well defined groups. Those attacking the mush-
rooms are called pathogens while those competing for the substrate are labeled indicators or
competitors. (Mushroom pathogens are either molds, bacteria, viruses or pests; indicators are
always fungi of some sort). In general, mushroom pathogens are not as numerous as the competitor
molds, though they can be much more devastating.

Not all molds and bacteria are damaging to the mushroom crop. To the contrary, several are
beneficial. These can not be called true “contaminants” since cultivators try to promote, not hinder,

236/The Mushroom Cultivator

Figure 176 High magnification scanning electron micrograph of Aspergillus spore be-
side germinating spore of Psilocybe cubensis.

their growth. To the beginner, however, they resemble real contaminants and therefore must be in-
cluded in this chapter. Examples of yield enhancing organisms are several thermophilic fungi and
bacteria, including:

Humicola

Torula

Actinomyces

Streptomyces

Select Pseudomonas and Bacillus species

These organisms are encouraged during the preparation of compost or during spawn run and are
rarely seen in agar or grain culture. Since they can not accurately be termed contaminants, the
aforementioned groups are not in the following key though they are fully discussed in the ensuing
descriptions.

Fungi, bacteria and viruses can be roughly delimited according to their size. All but viruses can
be detected by the home cultivator. Viruses can prevent fruiting, malform the mushroom fruitbody,
and expose the crop to further infestations from other pathogens. Since detecting viruses is beyond
the means of home cultivators, they have also been excluded from this key.

The Contaminants of Mushroom Culture/237

RELATIVE SIZES OF THE CONTAMINANT GROUPS

Size

Organism

(in microns)

Method of Defection

Viruses

.01 -.20

X-ray defraction, transmission electron microscopy and ultracentrifuge.
Typically attached to other larger partices, occurring within cells, or
are present in large conglomerate colonies. Often associated with
bacteria.

Bacteria

.40-5.0

Detected by electron microscopy, light microscopy and ultracentri-
fuge. Large colonies visible to unaided eye. Sometimes associated
with mushroom spores or mycelium.

Fungi

2.0-30.0

Detected by light microscopy. Large colonies visible to unaided eye.
Associated with a larger spore generating structure, often chain-like in
form.

Particulates screened out by
.3 Micron HEPA (High Efficiency

Particulate Air) Filter

45 microns
Visible to Eye,

Viruses (.003-. 05 microns)

Tobacco Smoke (. 1 -1 )

Bacteria (3-5 microns)

Fungus Spores (5-30 microns)

Plant Spores (10-80 micron:

Rain Droplet (600-10,000 microns)

Figure 177 Diagram illustrating comparative sizes of airborne particulates.

238/The Mushroom Cultivator

£2

What follows is a rudimentary key to the major contaminant groups encountered in mushroom
cultivation with the exception of insects and viruses which are discussed in later sections. Though
thousands of species of fungi exist in nature, only a small fraction are repeatedly seen in the course
of mushroom culture. Hence, this key is limited to that small sphere of microorganisms and does
not propose to be an all encompassing guide to the molds. Nevertheless, this key should prove to
be a valuable resource for anyone interested in improving their cultivation skills. Some contami-
nants are keyed out more than once if occurring in various habitats, or if exhibiting significant color
changes. Since color has some emphasis in this key and that feature can be substrate specific, the
authors presume the agar medium employed is 2% malt based, the spawn carrier is rye grain or
sawdust/bran, and the fruiting substrate is one outlined in this book.

Once led to a particular genus, refer to its description. If in doubt, a quick look under a medium
power (400 X) microscope should readily discern one contaminant from another. If the contami-
nant can be identified but its source can not, turn the chapter entitled Cultivation Problems and
Their Solutions. One or more of the common names have been listed under each competitor.
Good luck, be meticulous in your observations and strictly adhere to the recommended measures of
control.

Contaminants encompassed by this key:

Alternaria

Aspergillus

Bacillus

Botrytis

Chaetomium

Chrysosporium

Cladosporium

Coprinus

Dactylium

Epicoccum

Fusarium

Ceotrichum

Monilia Papulospora

Mucor Penicillium

Mycelia Sterilia Pseudomonas
Mycogone Rhizopus

Neurospora Scopulariopsis

Sepedonium

Trichoderma

Trichothecium

Verficillium

Yeasts

A KEY TO THE COMMON CONTAMINANTS IN MUSHROOM CULTURE

This key is easy to use. Simply follow the key lead that best descibes the contaminant at hand.
When the key terminates at a specific contaminant, turn to the descriptions immediately following
this key and then refer to the photographs and any related genus mentioned. To confirm the identity
of any contaminant, compare its sporulating structures with the accompanying microscopic illustra-

tions and/or micrographs.

la Contaminant parasitizing the mushroom fruitbody (a patho-
gen) 2

1 b Contaminant not parasitizing the mushroom fruitbody (an in-
dicator) 7

2a Contaminant causing mushrooms to become watery, slimy,
or to have lesions from which a liquid oozes but not covered

with a powdery or downy mycelium 3

2b Contaminant not as above but covering mushrooms with a
line powdery or mildew-like mycelium 4

The Contaminants of Mushroom Culture/239

3a Droplets forming across the cap and stem but lacking sunken
lesions. Mushrooms eventually reduced to a whitish foam-
like mass

3b Cap not as above but first having brownish spots that enlarge,
deepen, and in which a grayish brown slime forms. Mush-
rooms eventually disintegrate into a dark slimy, oozing mass

4a Contaminant eventually sporulating as a green mold on the
mushroom. Usually preceded by an outbreak of green mold
on the casing layer

4b Not as above

5a Contaminant appears on the casing soil as a fast running
grayish cobweb-like mycelium, enveloping mushrooms in its
path. (Spores usually three or more celled and 20 x 5
microns in size. If two celled, not acorn-shaped)

5b Contaminant attacking the mushroom but usually not ap-
pearing on the casing layer. (Spores single celled or if two
celled, resembling a roughened acorn and measuring much
less than above)

6a Contaminant turning young mushrooms into a rotting amor-
phous ball-like mass from which an amber fluid oozes upon
cutting. Stem typically not splitting or peeling. (Spores one
and two celled, the latter being darkly pigmented and acorn-
shaped) .

6b Contaminant afflicting young mushrooms as described
above but those parasitized not exuding amber fluid when cut
open. Stem in more mature mushrooms often splitting and
peeling, causing the mushrooms to tilt. (Spores one celled) .

Causal organism not known
“Weepers”

Pseudomonas tolassii
Bacterial Blotch
Bacterial Pit

Trichoderma viride
Trichoderma koningii
“Trichoderma Blotch”

Dactlyium dendroides
“Cobweb Mold”

Mycogone pernciosa
“Wet Bubble”

Verticillium malthousei
“Dry Bubble”

7a Contaminant in the form of another mushroom whose cap

deliquesces (melts) into a blackish liquid with age Coprinus spp.

“Inky Cap”

240/The Mushroom Cultivator

7b Contaminant not as above

8a Contaminant becoming pinkish to reddish to purplish col-
ored in age

8b Contaminant not as above

9a Occurring on compost or the casing layer

9b Occurring on nutrient agar media and on grain

1 0a Mycelium fast growing, aerial, and never having a frosty tex-
ture. Pinkish with spore maturity. (Spores unicellular with
nerve-like ridges longitudinally arranged and ellipsoid)

10b Mycelium slow growing, appressed, and developing a frosty
texture. Often becoming cherry red. (Spores cylindrical and
lacking nerve-like ridges)

11a Mycelial network of contaminant not well developed, not
clearly visible to the unaided eye, often slime-like

1 1 b Mycelial network of contaminant well defined and easily dis-
cernible to the naked eye, not slime like

12a More frequently seen in agar culture, (Spores produced by
simple budding, ovoid, single celled)

1 2b More frequently seen in grain culture. (Spores produced on a
short conidiophore, sickle shaped, and multicelled)

13a Mycelium fast growing and aerial. (Spores with nerve-like
ridges and ellipsoid)

1 3b Mycelium typically slow growing and appressed. (Spores two
celled, without ridges, and pear-shaped)

14a Contaminant slime-like in form

14b Contaminant mycelium-like or mold-like in form

15a Non-mofile (not moving spontaneously). Spores relatively
large, 4-20 microns in diameter. Not affected by bacterial an-
tibiotics such as gentamycin sulfate

10
1 1

Neurospora sp.
“Pink Mold”

Geotrichum “Lipstick Mold”

The Yeasts
see Cryptococcus

Fusarium

“Yellow Rain Mold”

Neurospora
“Pink Mold”

Trichothecium sp.

“Pink Mold”

The Yeasts

(see Cryptococcus and
Rhodotorula under Torula)

The Contaminants of Mushroom Culture/241

15b Motile (moving spontaneously). Spores relatively minute,
rarely exceeding 2 microns in diameter. Growth prevented
by bacterial antibiotics such as gentamycin sulfate .......

1 6a Cells rod-like in shape. Gram positive (retaining a violet dye
when fixed with crystal violet and an iodine solution)

1 6b Cells variable in shape. Gram negative (not retaining a violet
dye when fixed with crystal violet and an iodine solution) . . .

1 7a Contaminant mold greenish with spore maturity

1 7b Contaminant mold blackish with spore maturity

1 7c Contaminant mold brownish with spore maturity. .

17d Contaminant mold yellowish with spore maturity

1 7e Contaminant mold whitish with spore maturity

1 8a Forming small burrs and usually olive green in color. (Spores
lemon shaped, enveloped in a sac-like structure (a perithe-
cium)

18b Not as above

1 9a Molds typically blue-green in color. (Conidiophore diverging
at apex into multiple chains of lightly pigmented single celled
spores)

1 9b Molds typically true green to yellow green in color. (Condio-
phore swollen at apex and bulb-like (capitate), around which
multiple chains of lightly pigmented single celled spores ex-
tend)

1 9c Molds forest green in color. (Conidiophore easily disassem-
bling in wet mounts and difficult to observe under the micro-
scope. Spores single celled, lightly pigmented, and encased
in a mucous-like substance)

1 9d Molds blackish green in color. (Conidiophores branching
into few forks at whose ends darkly pigmented spores form,
often two celled.)

Bacillus
“Wet Spot”

Pseudomonas
“Bacterial Blotch”

25
28

Chaetomium olivaceum
“Olive Green Mold”

Penicillium spp.
“Blue Green Mold”

Aspergillus spp.
“Green Mold”

Trichoderma spp.
“Forest Green Mold”

Cladosporium spp.
“Blackish Green Mold”

242/The Mushroom Cultivator

20a Mold colony appressed, resembling a dark PeniciHium- like
mold, but not aerial

20b Mold colony aerial, not PeniciHium-like

21a (Spores elongated and ornamented with ridges, generaly ex-
ceeding 20 microns in length and 5 microns in diameter) . .

21b (Spores spherical, not ornamented with ridges, generally less
than 5 microns in diameter)

22a Most frequently seen on compost. Resembling black
whiskers. (Forming a conidiophore that diverges into multi-
ple stalks at whose ends are chains of darkly pigmented
spores)

22b Most frequently seen in agar and grain culture. Resembling a
forest of dark headed pins. (Forming a sporangiophore con-
sisting of single stalk at whose end a ball-like sporulating
structure is attached)

23a Conidiophore appearing swelled at apex; partially covered by
a sporulating membrane

23b Conidiophore not swelled as above; apex totally covered by
sporulating membrane

24a Mold developing small bead-like masses of cells (easily visi-
ble with a magnifying lens). Never producing cup-like fruit-
bodies. (Darkly pigmented cells clustered on a mycelial mat;
spores lacking)

24b Mold not developing the ball-like clusters of the above.
Sometimes producing cup-like fruitbodies. (Spores produced
in bunches in a grape-like fashion)

Alternaria spp.
“Black Mold”

Aspergillus spp.
“Black Mold”

Doratomyces stemonitis
“Black Whisker Mold”

Rhizopus

“Black Bread Mold”
“Black Pin Mold”

Mu cor

“Black Pin Mold”

Papulospora byssina
“Brown Plaster Mold”

Botrytis “Brown Mold”

The Contaminants of Mushroom Culture/243

25a Mold forming a corky layer between the casing layer and the
compost, and mat-like. (Spores borne on short vase shaped

pegs) Chrysosporium luteum

“Yellow Mat Disease”
“Confetti”

25b Mold not forming a corky layer and appearing mat-like.

(Spores not borne in the manner above) 26

26a Not occurring on compost. (Conidiophores short, arising
from cushion shaped cells. Spores, if reticulated, appear to

be composed of several tightly compacted cells) Epicoccum

“Yellow Mold”

26b Frequently seen on compost but not exclusively so. (Conidio-
phores not as above. Spores appearing unicellular) 27

27a Spores large, exceeding 5 microns in diameter, and of two
types. Some spherical and spiny, forming singly at the end of
individual hyphal branches; others vase shaped arising singly
or in loose clusters from an indistinct, hyphal-like conidio-

phore) . . . Sepedonium

“Yellow Mold”

27b Spores small, less than 5 microns in diameter, ovoid, form-
ing on chains arising from a head-like structure positioned at

the apex of a long stalk Aspergillus spp.

“Yellow Mold”

28a Appearing as a dense plaster-like or stroma-like mycelium.

(Condiophore brush shaped (pencillate)) Scopulariopsis

“White Plaster Mold”

28b Mycelium not plaster-like. (Conidiophore not brush shaped
(pencillate)) 29

29a Spores forming from hyphae in chains Monilia

“White Flour Mold”

29b Spores absent, not forming from hyphae Mycelia Sterilia

(see also: Mucor and
Sepedonium).

244/The Mushroom Cultivator

VIRUS

Common Name: Die-back Disease; La
France Disease, Mummy.

Habitat and Frequency of Occurence:

An infrequent and difficult to detect disease.
Their habitats are other larger particles or or-
ganisms.

Medium through which contamination
is spread: Primarily from infected mycelium
or from the spores of diseased mushrooms.
Dieleman-van Zaayen (1972) found that the
most common way virus spreads is through
the anastomosing (“merging”) of healthy
mycelia with infected mycelia that was left-
over from previous crops. Once anasto-
mosed, the virus particles spread throughout
the mycelial network of the new mycelium.

Measures of Control: Thorough disinfection of the growing room between crop rotations by
steam heating for 12 hours at 158-160° F.; the installation of high efficiency spore filters to screen
particulates exiting the growing environment; the disinfection of floors and hallways leading to and
from the growing room with 2% chlorine solution; and picking diseased mushrooms while the veil
is intact before spores have the opportunity to spred. Isolation of infected crops from adjacent rooms
or those newly spawned helps retard the spread of this disease. Other measures of control include
the placement of disinfectant floor mats to prevent the tracking in of virus-carrying particles on work-
er’s shoes and the maintenance of strict hygienic practices at all times, particularly between crops.

Macroscopic Appearance: On nutrient agar media, infected mycelia slows or nearly abates in
its rate of growth as the disease progresses throughout the mycelial network. When running
through the casing layer, large zones one to three feet in diameter remain uncolonized. In some
cases the mycelia, once present, disappears from the surface. Fruitbodies may not form at all, or
when they do, the mushrooms are typically deformed (dwarfed or aborted), often with watery or
splitting stems, and brown rot. The caps prematurely expand to plane. Virus infected cultures can
exhibit any combination of the above described symptoms.

Microscopic Characteristics: Particles typically ovoid to polyhedral, measuring 25 or 34 nano-
meters. Elongated particles measure 1 9-50 nanometers. Virus particles dwell within hyphal cells or

The Contaminants of Mushroom Culture/245

on the surfaces of spores. They are detectable only through transmission electron microscopy or
ultracentrifuging.

History, Use and/ or Medical Implications: Responsible for many plant, animal and human dis-
eases. Typically viruses are associated with larger carrier particles, particularly bacteria.

Comments: Virus is most likely introduced during or directly after spawning. Infected farms experi-
ence losses up to 70%. First reported from Europe, measures of control and prevention have been
developed and successfully tested by the Dutch. Most notably, virus spreads by attaching itself to
mushroom spores which then become airborne. Virus also spreads through the contact of healthy
mycelia with diseased mycelia. Afflicted mushrooms are soon exploited by a host of other parasites,
making a late and accurate diagnosis of this contaminant difficult.

Undoubtedly, virus is the cause of what many have noted as “strain degeneration”. Heat treat-
ment of infected strains grown on enriched agar media at 95° F. for three weeks has been sug-
gested as one remedy for curing diseased mycelia. (See Gandy and Hollings, 1 962 and Rasmussen
et al„ 1972).

Van Zaayen (1979) and others have noted that Agaricus bitorquis seems resistant to virus dis-
ease even when inoculated with in vitro particles. Another species of Agaricus, called Agaricus
arvensis, exhibits similar virus resistant qualifies.

Virus-like particles have also been found in Lentinus edodes by Mori et alia (1 979) but do not
adversely affect fruitbody formation or development. These same researchers reported that this spe-
cies’ viruses can not be transmitted to other mushrooms or plants, a fact they attributed to the inter-
feron producing properties of the shiitake mushroom. No work with infected strains of Psilocybe are
known. Only a fraction of wild mushrooms harbor virus-like particles.

246/The Mushroom Cultivator

ACTINOMYCES

Class: Actinomyces
Order: Actinomycetales
Family: Actinomycetaceae
Common Name: Firefang.

Greek Root: From “actino” meaning rayed
or star-like and “myces” or fungus, in refer-
ence to its characteristic appearance when
colonizing straw or straw/manure compost.
Habitat & Frequency of Occurrence:
Many species thermophilic; thriving in the
115-1 35 °F. temperature range and com-
monly found in decomposing straw, horse
and cow manures. Actinomyces are impor-
tant soil constituents. They thrive in aerobic,
well prepared mushroom composts.
Medium Through Which Contamina-
tion Is Spread: Primarily air; secondarily the
straw used in compost preparation.

Measures of Control: Generally no controls are necessary during compost preparation. However,
Actinomyces can cause spontaneous combustion in wet, compacted straw. Covering stored baled
straw from excess water absorption should be adequate protection from Actinomyces and the ther-
mogenic reactions they cause.

Macroscopic Appearance: Grayish to whitish speckled colonies, readily apparent on dark com-
posted straw.

Microscopic Characteristics: Composed of an extensive, fine hyphal network that rarely
branches. Rod-like spores form when the filaments break at the cell wall junctions. The filamentous
hyphae and spores are minute, measuring only 1 micron in diameter. Within each cell, no well de-
fined nucleus is discernible. Lacking differentiated spore-producing bodies, Actinomyces are Gram-
positive.

History, Use, and/or Medical Implications: Few species pathogenic. Amongst agricultural
workers in the same position, males are three times more susceptible to this bacterium than females
(see Cruickshank et al . , 1 973). Two notable species causing serious diseases (actinomycosis) of the
skin and oral cavity in humans are Actinomyces bovis and Actinomyces israelii. Generally, these

Figure 178 Drawing of Actinomyces.

The Contaminants of Mushroom Culture/247

species behave as secondary infectious organisms. Penicillin is often used for treatment. Actino-
mycin, a potent antibiotic compound interfering with RNA synthesis, is derived from this group of
bacteria.

Although the likelihood of mushroom growers contracting actinomycosis is remote, workers
spawning compost are exposed to high concentrations of Actinomyces spores and often report less
severe, temporary allergic reactions. Therefore, the use of a filter mask when spawning large
volumes of compost is advisable.

Comments: The Actinomyces resemble both bacteria and fungi and have alternately been called
one or the other. Presently, the prevailing belief is that they are filamentous (Gram-positive) bacteria
because they are prokaryotic (lacking a defined nucleus), are inhibited by bacterial antibiotics and
not affected by fungal antibiotics;, and lack the chitin-like compounds so typical of the true fungi.
The hyphal filaments of Actinomyces are one fifth to one tenth as thick as those of true fungi.

Actinomyces are commonly called Firefang for their ability to cause spontaneous combustion
of decomposing materials. (Spontaneous combustion is prevented by proper composting
practices.) Many of these bacteria/fungi are true thermophiles and can live aerobically or anaerobic-
ally. Actinomyces is the major microorganism selected to colonize the compost during Phase II.
When the finished compost is spawned, Actinomyces are consumed by the mushroom mycelia.

See also Streptomyces. See Color Photo VIII.

BACILLUS

® 0

I e

%r *
% Do

Bituffuj

I igure 179 Drawing of endospore forming
Bacillus cells as they appear through a micro-
scope and without special stains.

Class: Schizomycetes

Order: Eubacteriales

Family: Bacillaceae

Common Name: Wet Spot; Sour Rot.

Latin Root: From “bacilliformis” meaning

rod-like, in reference to its characteristic

shape.

Habitat & Frequency of Occurrence: Liv-
ing within a broad range of habitats. Bacillus
grows on almost anything organic that is
moist and is surrounded by oxygen. It is par-
ticularly common in soils.

Medium Through Which Contamina-
tion Is Spread: Primarily through the air;
secondarily through water, grain, soils, com-
posts, insects, tools and workers.

Measures of Control: Air filtration through high efficiency particulate air filters; thorough steriliza-
tion of grain; and proper storage and use of relatively “clean” grains. The addition of antibiotics to
agar media (gentamycin sulfate, penicillin, streptomycin, aureomycin, etc.) hinders or prevents the
growth of these contaminants. Endospores are neutralized by exposure to moist heat, such as the
steam generated within a pressure cooker at temperatures of 250 °F. an^gjl5 psi pressure for a full
hour Temperatures as low as 140°F. kill the vegetative parent celM}i0§iot the endospores they
form. \Sj g j

Macroscopic Appearance: A dull gray to mucus-like brownish ^Wnelftnaracterized by a strong
but foul odor variously described as smelling like rotting apples, dirty socks or burnt bacon. Bacillus
makes uncolonized grain appear excessively wet, hence the name Wet Spot . Pallid to whitish
ridges along the margins of individual grain kernels characterize this contaminant.

Microscopic Characteristics: Rod-like or cylindrical in shape, measuring 0.2-1 .2 microns in di-
ameter and 1 -5 microns in length. When wet mounts are viewed through a microscope, Bacilli ex-
citedly wriggle back and forth. Species move by the vibrating action of flagella ( hairs ) that outline
each cell. These flagella are difficult to observe microscopically without using specific staining tech-
niques. Bacilli are encapsulated by a thin but firm slime and conglomerations of cells give infected

... , -T^^rr mt m

The Contaminants of Mushroom Culture/249

Figure 1 80 Bacillus, the Wet Spot bacter-
ium, as it appears on grain.

Figure 181 Scanning electron micro-
graph of rod shaped bacteria on a spore of
Panaeolus acuminatus.

Figure 182 Scanning electron micro-
graph of rod shaped bacteria on mycelium
of Psilocybe cubensis.

250/The Mushroom Cultivator

grain a slimy appearance. Bacillus primarily reproduces through simple ceil division. In times of
adverse environmental conditions, especially heat, a single hardened spore forms within each par-
ent cell body. These endospores show an extraordinary resistance to heat, are low in water content
and are unaffected by drying. Species in this genus are Gram positive.

History, Use, and/or Medical Implications: The most notable species in the genus is Bacillus
anthracis, the cause of the hideous Anthrax disease that killed several thousand sheep when an
United States Army experiment went awry in Utah during the 1950s. Home cultivators are, how-
ever, unlikely to be exposed to this species. Most endospore forming bacteria are not virulent.
Bacillus subtilis, the bacterium spoiling grain spawn, is being developed to replace E. coli as a re-
combinant-DNA fermentor. Clostridium is a genus similar to Bacillus except that it is anaerobic.
That genus is reknowned for one toxic species in particular: C. botulinum, the cause of botulism.
Comments: A pernicious and tenacious competitor, Bacillus contamination is the most difficult to
control. At room temperature, a single cell reproduces every 20 minutes and will multiply into near-
ly a million daughter bacteria in only seven hours. In another seven hours each one of those million
bacteria divide into a million more cells. Thus, in less than fourteen hours, one trillion bacteria
evolve from a single parent cell!

The phenomenonal reproductive capability of Bacillus and other bacteria poses a formidable
threat to the spawn maker. Although parent cells are easily destroyed, their endospores are not.
Under dry conditions, endospores form in increasing numbers as temperatures rise to 130° F. In
boiling water (212° F.), endospore viability markedly decreases. (Ninety percent of Bacillus spores
are killed in only one minute at 21 2 ° F.). At the higher temperatures and pressures within an auto-
clave the survivability of Bacillus spores falls well below 1 %. Nevertheless, this 1 % seriously ob-
structs any attempt at grain culture given Bacillus’ rapid reproductive capability. This problem is
compounded if the bacteria count in the grain is initially high.

In one study (Shull and Ernst, 1962), the thermal death time (TDT) of an exposed Bacillus
stearothermophilus population of 1,300,000 endospores was pinpointed at 250 °F. for 13 min-
utes. (In a pressure cooker at sea level, 250 °F. corresponds to 15 psi). Food researchers con-
cerned with food-spoiling bacteria (particularly Clostridium) have shown that endospore endurance
to heat is directly related to the amount of calcium in the host substrate. Once formed, endospores
can sit dormant for extended periods of time. Even endospores removed from the stomachs of
mummies have proved viable after hundreds of years.

Although an autoclave may read a certain temperature, the grain within the spawn containers
may be well below that reading. To guarantee adequate steam penetration, the water in pressure
cookers should be brought to a boil for 5 minutes before closing the vent valve. Furthermore, bacte-
ria within the spawn container are partially protected from the sterilizing influence of steam by the
structural cavities of the grain medium. This delay in steam penetration time is especially character-
istic of large, heavily packed autoclaves.

Despite the fact that autoclaving for one hour at 15 psi is sufficient to kill most contaminants,
grain having initially high bacteria populations may require sterilization at higher temperatures and
for prolonged periods of time. Autoclaving quart jars for 1 hour at 270 ° F. (which is equivalent to

The Contaminants of Mushroom Culture /251

27 psi) is sufficient to neutralize grain heavily infested with endospore forming bacteria. If convert-
ing a standard home pressure cooker for this purpose, contact the manufacturer about stress limita-
tions and follow all safety recommendations.

If “sterilized” rye grain spontaneously contaminates with bacteria before inoculation and the
grain is the cause, it is best to replace the grain with a cleaner one than to undergo the expense and
time of double sterilization. Some spawn laboratories regularly precook their grain for approximate-
ly 2 hours in water at a low boil. Excess water is allowed to drain from the grains which are then
placed into the spawn container and sterilized at standard time and pressure.

The most practical method for eliminating bacterial endospores involves soaking the grain at
room temperature 24 hours prior to sterilization. Endospores, if viable, will germinate within that
time frame and then be susceptible to standard sterilization procedures. And, new endospores won’t
form in the moist environment of the resting jar of grain.

Bacillus subtilis var. mucoides is the common bacterium responsible for spoiling spawn
media. If allowed to proliferate, this contaminant wreaks havoc in a spawn laboratory, necessitating
a complete shut-down of operations. Spores and even strains of mushroom mycelium can become
hosts for Bacillus, carrying bacteria on their hyphae (see Figs. 1 81 & 1 82), and then contaminating
any media onto which the mushroom mycelia is transferred.

Many bacteria are rod-shaped and the term bacillus has been loosely used to describe them.
The genus concept of Bacillus, however, has been narrowed considerably with time; Bacillus is
now defined as Gram positive rod-like, aerobic bacteria that form spores.

According to Park and Agnihotri (1969), Bacillus megaterium stimulates primordia formation
in certain strains of Agaricus brunnescens (bisporus). (See Appendix II for a futher discussion on
the influence of bacteria on fruiting). Another species, Bacillus thermofibricolous, if introduced at
spawning, inhibits the growth of competitor molds in rice bran/ sawdust spawn prepared for shiitake
cultivation according to Steineck (1973).

See also Botrytis and Scopulariopsis.

300/The Mushroom Cultivator

PEN I Cl ILIUM

Class: Fungi Imperfecti
Order: Moniliales
Family: Eurotiaceae

Common Name: The Bluish Green Mold.

Latin Root: From “penicillum” meaning a
brush-like tuft of hairs, so named in reference
to the shape of the sporulating body.
Habitat & Frequency of Occurrence: An
extremely common contaminant. Although
not as prevalent in nature as Cladosporium,
Penicillium is the most prevalent of indoor
contaminants, a fact that is undoubtedly re-
lated to human eating habits. Penicillium
species abound on foodstuffs such as fruits,
cheeses and stored grains. Many species pre-
fer habitats with an acid pH. Penicillia are oc-
casional to frequent on under-developed
mushroom compost, casing soil and on dis-
carded mushroom debris.

Medium Through Which Contamination Is Spread: Primarily through the air, although stored
grain and other foodstuffs, as well as humans are the most frequent carriers of this mold.

Measures of Control: Air filtration; removal of waste products; isolation of contaminated cultures;
and maintenance of a high level of hygiene.

Macroscopic Appearance: Appearing as a granular or powdery bluish green mold, often with a
broad whitish rim of new growth. Some species, less frequently encountered, are whitish, yellowish
or even reddish in color. Many species exude droplets of fluid from their surfaces having antibiotic
properties.

Microscopic Characteristics: Conidiophores arising singly, long, and branching near the apex
into short chains of globose, green, dry conidia. Compared to mushroom spores, the conidia of
Penicillia are minute, measuring only 2-4 microns in diameter.

History, Use and/or Medical Implications: In 1928-1929 while Dr. Alexander Fleming was
studying Staphylococcus aureus, he noticed that a green mold contaminant inhibited his cultured

Figure 217 Drawing of sporulating struc-
ture characteristic of Penicillium molds.

The Contaminants of Mushroom Culture/301

Figure 218 Scanning electron
micrograph of Penicillium.

bacteria when the two grew in close proximity. A fluid that was being exuded from the fungus
caught his curiosity. Upon reporting his finding, collegues later found the fluid contained a powerful
new antibiotic which was named penicillin. He had, in fact, cultured Penicillium notatum Westl.
Currently penicillin is corqhn^lially produced by high yielding strains of Penicillium chrysogenum
Thom. Through its useL®ii'8f|s of people have been cured of illnesses that were previously untreat-
able. From the widespfl^yi&J and abuse of this drug, however, new, more virulent and penicillin
resistant strains of bacteria h^ve evolved.

From the production of steroids to the making of roquefort cheese (by Penicillium roquefortii),
this genus is resplendent with species of proven value to man. Few, if any, are pathogenic.

Comments: Since the high count of Penicillium spores indoors is directly traceable to decompos-
ing foodstuffs, one can reduce the prevalance of this contaminant by simply following good hygien-
ic practices. Penicillium, a prolific spore producer, is an ubiquitous fungus. It is probably the most
common contaminant seen in the laboratory. Although Penicillium can attack compost, casing soil
and mushroom debris, it is not as prevalent as Trichoderma in these habitats. Other green molds,
similar in appearance, are Cladosporium and Aspergillus.

Penicillium sometimes contamine'es poorly prepared compost or spawned compost that has
undergone secondary heating. Here, grain kernels formerly colonized by mushroom mycelium be-
come susceptible to weed molds such as Penicillium and Doratomyces and then spread onto the
compost and/or casing soil.

Differing from Aspergillus and Trichoderma in the shape of the conidiophore.

For further information:

“The Penicillia” by Raper and T"om (1949) who recognized 1 38 species at the time of pub-
lication.

See Color Photograph 20.

302/The Mushroom Cultivator

UZUZSi

RHIZOPUS

Class: Zygomycetes
Order: Mucorales
Family: Mucoraceae

Common Names: Bread Mold; The Pin
Mold.

Latin Root: From the prefix “rhizo”, per-
taining to roots, and the suffix “pus” or foot,
in reference to the rhizoids at the base of the
sporangiophore that is characteristic of some
species in this genus.

Habitat & Frequency of Occurrence: A

saprophyte, commonly seen in both agar and
grain culture. Rhizopus naturally inhabits
dung and soils and is a decomposer of dead
Figure 219 Drawing of asexual sporangio- plant and animal matter. Within the home,

phore and sexual zygosporium of Rhizopus. this contaminant is most often seen on old

bread or on poorly stored grain and fruits.

Medium Through Which Contamination Is Spread: Primarily air.

Measures of Control: Air filtration; strict adherence to general hygienic practices; and steam steri-
lization of grain and agar media.

Macroscopic Appearance: Similar to Mucor. When sporulating, Rhizopus appears as a a dense
mat of tall, aerial, vertically oriented hyphae upon which sit dark grey to grey black heads. It resem-
bles a forest of pins.

Microscopic Characteristics: A creeping hyphal network that gives rise to individual, vertically
oriented stalks that are unbranched and at whose base distinct rhizoids can be attached. The apex is
swelled into a vesicle upon which a dark spherical body (sporangium) rests. This sporangium does
not fully envelope the sporangiophore. Hence, the sporangiophore swells before contacting the
sporangium. The sporangium is a mass of spores within a thin envelope of tissue that soon disinte-
grates and frees the asexual spores. Joining these individual sporangiophores are long interconnect-
ing mycelial veins called stolons. Mating can also occur betwen two sexually complementary
hyphae and results in the formation of a globose reproductive body, a zygosporium. (See Fig. 219).
Its mycelia lacks distinct cell walls.

The Contaminants of Mushroom Culfure/303

History, Use and/or Medical Implications: In itself, not a pathogen to man. Reports in the
medical literature have in the past blamed Rhizopus for zygomycosis when in fact other related
genera— Absidia and Mucor— were responsible.

Rhizopus stolonifer, the black bread mold, is also utilized in the commerical production of fu-
maric acid and cortisone. Other species in the genus secrete assorted alcohols and acids as meta-
bolic waste products.

Comments: Along with Aspergillus and Penicillium, species of this genus are the primary con-
taminants of grain spawn. Rhizopus is very rapid growing, and is called the Pin Mold for the shape
of the spore producing body. Rhizopus stolonifer (= Rhizopus nigracans) is called the Black Pin
Mold and can elevate the substrate temperature from room temperature to the 95-104° F. range.
At this level, the populations of the true thermophiles increase dramatically, further heating up the
host substrate to temperatures lethal to the mushroom mycelium.

See also Mucor, a mold that is closely related to Rhizopus, but whose sporangium completely
covers the apex of the sporagiophore.

Figure 220 Rhizopus, the Black Pin Mold, on malt agar.

304/The Mushroom Cultivator

Figure 221 Drawing of sporulating struc-
ture typical of Scopulariopsis.

SCOPULARIOPSIS

Class: Fungi Imperfecti
Order: Moniliales
Series: Annelosporae
Common Names: White Plaster Mold;
Flour Mold.

Latin Root: From “scopulatus” meaning
broom-like or brush shaped, in reference to
the structure of the sporulating reproductive
body.

Habitat & Frequency of Occurrence: A

saprophyte, occasionally seen in composts
that have been over-watered or are too high
in nitrogen. Scopulariopsis also forms on the
casing during the fruiting cycle. It naturally
grows in soils, on hay, on rotting leaves and
on other decaying plant material including
grain. This group of molds generally prefer
an alkaline pH.

Medium Through Which Contamination Is Spread: Primarily airborne spores, spent compost
and insects; and from materials previously in contact with this contaminant that were not thoroughly
cleaned before use.

Measures of Control: Proper preparation and sufficient air during Phase II composting discour-
ages this fungus. Atkins (1974) reported that excessive moisture and subsequent anaerobic
pasteurization were the two main factors contributing to the spread of the White Plaster Mold. Be-
fore filling, the addition of gypsum to an overly wet compost will bind loose water, a condition
favorable to this mold.

Macroscopic Appearance: Circular colonies of densely matted, whitish mycelia; with age devel-
oping slight pinkish tones. This mold often appears as “splotches”, mostly on the compost bed and
to a lesser degree on the casing soil.

Microscopic Characteristics: Conidiophores short, soon branching, delineating into several
elongated cells which then give rise to short chains of globose, hyaline, finely warted, dry conidia
that measure 5-8 x 5-7 microns. Annular zonations are present at the junction of the sporogenous
cells and the first spore in the conidial chain. Terminal cells in the chain are the oldest and typically

S2VL3M

isessa

The Contaminants of Mushroom Culture/305

the largest. The conidiophores generally resemble that of Penicillium and thus are described as
pencillate, or brush shaped.

History, Use and/or Medical Implications: One species toxic to humans: Scopulariopsis
brevicaulis (Saccardo) Brainer. This species usually attacks tissue already diseased by other micro-
organisms. It is an improbable threat to the health of mushroom cultivators.

Comments: Scopulariopsis fimicola is the White Plaster Mold seen on compost beds. It is very de-
trimental to the growth of mushroom mycelia. Its presence is usually an indication of a short, wet
and over-mature compost. This condition predisposes the compost to a difficult Phase II with dense
anaerobic areas, ammonia-lock and consequently high pH levels. All these factors contribute to the
growth and spread of Scopulariopsis fimicola, the species of White Plaster Mold most frequently
seen in mushroom culture.

Contamination can also arise from within the mushroom house if there has been a prior history
of problems with this contaminant and if strict contamination control procedures have not been in-
stigated. Not surprisingly, one often finds Scopulariopsis with the Inky Cap (a Coprinus species)
which is also associated with residual ammonia in composts.

See also Papulospora (P. byssina Hots.), a genus containing the Brown Plaster Mold whose
early stages of growth resemble the White Plaster Mold.

306/The Mushroom Cultivator

SEPEDONSUM

Class: Fungi Imperfecti
Order: Moniliales
Series: Aleurisporae

Common Names: Yellow Mold; White
Mold.

Habitat and Frequency of Occurrence:
Occasionally to frequently encountered on
agar; more common on compost; and para-
sitic on wild mushrooms (both Basidiomy-
cetes and Ascomycetes).

Medium Through Which Contamina-
tion Is Spread: Primarily through the air,
Figure 222 Drawing of sporulating struc- but also from spent compost,
ture and flask shaped conidia. Methods of Control: Air filtration; strict

maintenance of hygiene in the laboratory and
growing room; the expeditious removal of spent compost; and the thorough disinfection of wooden
compost containers.

Macroscopic Appearance: On malt agar and on rye grain appearing as a fast growing whitish
mold, very similar to cottony mushroom mycelia and frequently mistaken for it. On compost it is a
fine white mold which with age becomes yellowish to golden yellow from spore production, it is not
as prolific a spore producer as the powdery Trichoderma. If spores are not produced at all, the my-
celia remains whitish. This mold attacks composts that otherwise have been properly prepared for
mushroom growing.

Microscopic Characteristics: Two types of spores formed. The more obvious are large, globose
chlamydospores ornamented with short spines and similar to those of Mycogone; except in this
genus a hemispheric foot cell, shaped like a teacup is absent. Conidiophores are simple, relatively
undeveloped, resembling mushroom mycelium and not easily distinguished from it except that they
lack clamps. Globose to vase shaped conidia develop terminally at the end of these branches, either
singly or in loose clusters.

The Contaminants of Mushroom Culture/307

History, Use and/or Medical Implications: Some species possibly toxic. It has been suggested
that this mold secretes a sweet odor nauseous to some mushroom workers and possibly the cause
of a little understood respiratory illness. Not much is known.

Comments: Sepedonium spores are noted for their heat resistance. It is a whitish mold until the
yellow conidia are produced. On malt agar media, Sepedonium is fast running, and out-grows most
mushroom mycelia. When the two fungi grow within close proximity to one another, a line of inhibi-
tion usually develops between the two. If conidiospores or chlamydospores are not produced, this
mold is difficult to identify. The conidiophores are indistinct, very much resembling its own myceli-
um. From the authors’ experience this contaminant is a vigorous competitor on agar media. Its ap-
pearance necessitates a thorough cleaning of the laboratory and spawn incubation environment. If
this mold contaminates grain spawn and goes undetected, use of this spawn in subequent inocula-
tions would be disastrous.

The second site of contamination is horse manure/ straw compost where it most frequently ap-
pears during spawn run. Only detrimental when iarge outbreaks occur, Sepedonium’s presence on
compost can be traced to insufficient pasteurization or spent compost residues in the trays or
shelves. Although not regarded as a serious competitor on mushroom compost, Sepedonium is an-
other fungus believed to be a food source for mites (Kneebone, 1961).

Sepedonium, like Mycogone, is an imperfect state of Hypomyces, a common parasite on
mushrooms. In the wild, Sepedonium chrysosperma parasitizes Boletus species (particularly B.
chrysenteron) and causes them to abort. The chlamydospores of Sepedonium are generally similar
to Mycogone.

See also Mucor and Mycelia Sterilia, two fast running whitish molds on agar media.

Figure 223 Sepedonium mold compet-
ing with Psilocybe cubensis mycelia on
malt agar media.

308/The Mushroom Cultivator

Figure 224 Drawing of the sporulating
structure of Torula, the Black Yeast

TORULA

Class: Fungi Imperfecti
Order: Moniliales
Family: Dematiaceae

Common Name: Black Yeast (Torula
nigra).

Latin root: From the same root as the adjec-
tival “torulosus”, meaning cylindrical shaped
with bulges and constrictions at regular inter-
vals, chain-like.

Habitat and Frequency of Occurrence:

Saprophytic, common. Many thermophilic
species participate in the decomposition of
straw and manure in the making of mush-
room composts. Although Torula is rarely
seen in agar culture, its cousin Rhodotorula,
a red yeast, is frequently seen.

Medium Through Which Contamination Is Spread: Primarily an airborne contaminant; sec-
ondarily transmitted through compost.

Methods of Control: None generally needed or desired. Torula is a beneficial, thermophilic
microorganism thriving in the 115-125° F. range.

Macroscopic Appearance: Whitish at first, then grayish, soon dark brown or jet black with spore
production. As Torula matures, the mycelium becomes covered with a mass of spores that give it a
soot-like appearance. On compost, this fungus appears similar to Humicola.

Microscopic Characteristics: Mycelium colorless or slightly pigmented. True conidiophores are
lacking. Hyphae abruptly terminate into conidia which are ovoid, translucent, dark brown, smooth
and produced in branched or unbranched chains by either of two methods. In one form, the more
mature spores of a conidial chain develop apicaily, with the younger spores arising from the spore
closest to the hyphal branch. (This is called basipetal development). In a second form, conidia can
develop by simple budding from the tips of a hypha, in a yeast-like fashion. The budding hypha nar-
rows towards the apex into immature spores and finally terminates with an attenuating tail. Freed are
conidia found singly or attached several at a time.

The Contaminants of Mushroom Culture/309

History, Use and/or Medical Implications: Not thought to be pathogenic. Confusion with
Cryptococcus has in the past given Torula an undeserved pathogenic reputation. Cryptococcosis in
the medical literature is often though incorrectly termed torulosis.

Comments: Torula, like Humicola, is an ally to the mushroom compost maker, converting am-
monic nitrogen into protein usable to the mushroom. Torula thermophila Cooney & Emerson is
the species most frequently seen in composting straw and manure. Originally isolated from chicken
droppings, this species is a true thermophile with a temperature range from 73-1 36 0 F., and an op-
timum of 1 04 0 F. The Torula genus is known for a number of thermophilic species that survive the
pasteurization process and flourish at standard Phase II conditioning temperatures (1 18-125° F).
When pasteurized compost is cooled down to room temperature, this fungus is rendered inactive
and in turn becomes a food source for the mushroom mycelium.

Rhodotorula reproduces very similarly to Torula. It is known as the Red Yeast, commonly con-
taminating agar cultures. Rhodotorula glutinis, a common soil inhabitant, may play an important
role in the reproductive cycle of the common Chantarelle mushroom, Cantharellus cibarius. Pure
cultures of Chantarelles have been difficult to obtain from wild specimens. And, Chantarelle spores
do not germinate using standard laboratory techniques. In 1979, a Sweedish mycologist named
Nils Fries discovered that, in the presence of Rhodotorula glutinis and activated charcoal, C.
cibarius spores readily germinate. Pure cultures of Chantarelles, once nearly impossible to obtain,
are now feasible. Other related yeasts may have a similar stimulatory effect on various mushrooms
species currently not prone to cultivation.

Torula species, as with most yeasts, are separated from one another largely by chemical

means.

310/The Mushroom Cultivator

TRICHODERMA

Class: Fungi Imperfecti
Order: Moniliales
Family: Moniliaceae

Common Names: Forest Green Mold;
Green Mold; and Trichoderma Blotch.
Greek Root: From “trichos” meaning hairy
and “derma” or skin.

Habitat & Frequency of Occurrence:

Very common on compost, casing soil and
to a lesser degree on grain and agar.
Trichoderma often parasitizes mushrooms
under cultivation and can inhibit or reduce
fruitings. Many species grow on wood or
Figure 225 Drawing of conidia and sporu- woody tissue and are abundant in peat,
lating structure typical of Trichoderma. Trichoderma frequently grows on the wood-

en trays holding compost.

Medium Through Which Contamination Is Spread: Primarily an airborne contaminant when
contaminating agar or grain cultures. On casing soils, it is introduced through the peat or humus.
Trichoderma is often spread during harvesting, bed cleaning or watering. Species in this genus
generally prefer an acid pH in the 4-5.5 (6) range.

Measures of Control: Careful picking; disposal of dead and diseased mushrooms; lowering of hu-
midity levels; lowering carbon dioxide and increasing air circulation to eliminate dead air pockets.
Use of clean casing materials lacking undecomposed woody tissue lessen the chance of Trichoder-
ma contamination. Isolated outbreaks of Trichoderma can easily be contained by one of several
methods. Since Trichoderma thrives in acid habitats, raising the pH of the surrounding soil inhibits
further growth. Perhaps the simplest way to raise pH is to cover the infecting colony with salt, sodi-
um hypochlorite or sodium bicarbonate (baking soda) or a solution thereof. Recognizing and
treating this fungus in its earliest stages, before many spores are produced, greatly reduces the risk
of satellite colonies spreading throughout the growing room. Mushrooms afflicted with Trichoderma
should be carefully isolated. All items coming in contact with it (tools, workers, etc.) should be
resanitized. Steam pasteurization at 160°F. for one hour effectively kills the spores of this fungus

The Contaminants of Mushroom Culture/31 1

Macroscopic Appearance: A cottony mold, growing in circular colonies on the casing soil or on
compost; grayish and diffuse at first; rapidly growing; and soon forest green from spore production.
On malt agar colonies of Trichoderma have an aerial, cottony and brilliant forest green mycelium
whereas Penicillium has an appressed, granular and blue green mycelium. Some infrequently en-
countered species are whitish or yellowish, but the majority of those seen in mushroom culture are
greenish shaded.

Parasitized mushrooms have dry brownish blotches or sunken lesions on the cap or stem.
They are often enveloped by a fine downy mildew that may eventually become greenish from spore
production, and are grossly misproportioned.

Microscopic Characteristics: Conidiophores clear, profusely branched upon whose ends small
bunches of ovoid greenish pigmented, smooth spores are borne. In many species uniquely shaped
sporogenous cells are present roughly resembling bowling pins and arranged as triads. After
squashing a sample for viewing under the microscope, the conidiophores readily disassemble and
are difficult to recognize. The freed conidia, however, are not arranged in linear chains as common-
ly seen in Aspergillus and Penicillium, but are in loose clusters or are scattered as individuals. A

Figure 226 Trichoderma-Wke mold parasitizing Psilocybe
cubensis.

312/The Mushroom Cultivator

most distinctive feature is that the conidia are encased in a mucus-like substance, making the spores
sticky. Spores measure 3-5 x 3-4 microns.

History, Use and/or Medical Implications: Not known to be pathogenic. One industrial ap-
plication utilizes Trichoderma, Pencillium and Cladosporium to precipitate precious metals such as
gold and platinum from solutions. The process is being patented.

Comments: In cased grain culture, Trichoderma is the most frequently encountered contaminant
on the casing layer and usually originates there. Upon casing, spores harbored in the peat infect ex-
posed grain kernels and sporulate. The contaminated kernels become a platform for further con-
tamination. The mold then spreads through the casing layer until it breaks through to the surface of
the casing layer. Also, Trichoderma is prone to casings with undecomposed woody tissue and
those incorporating potting soils. Trichoderma is also caused by excessively wet casings applied to
sterile grain spawn.

Trichoderma is an ubiquitous fungus that is encouraged by improperly adjusted environmental
parameters. Conditions of excessively high and prolonged humidity in combination with stagnant
air and high carbon dioxide levels tip the ecological balance of the casing soil’s micro-ecology in fa-
vor of this contaminant. Once Trichoderma populations bloom, this mold quickly infects newly
formed primordia and developing fruitbodies which become deformed. This pathogen also grows
on discarded mushroom debris, particularly stem butts.

Afflicted mushrooms have brownish specks or lesions on the stem, especially near the base or
apex. A fuzzy mycelium similar to Verticillium may be present on the cap. These lesions are dry,
whereas the blotches caused by bacteria tejid to be moist. The growth of the fruitbody is abruptly ar-
rested by this mold. Under extreme eqViffljfons this mold sporulates directly on the mushroom, be-
coming green in color. Adjacent rti^M|ns, newly formed pinheads and subsequent crops need
not be affected if air circulation is^^a^d to proper levels and if humidity is decreased to within
tolerable limits (3-5 exchanges of air per hour while maintaing 85-92% humidity). Trichoderma is
alleged to secrete toxins that inhibit mushroom primordia formation and growth.

Another problem with Trichoderma is that its spores are utilized by red pigmy mites as food.
Trichoderma spores are sticky and attach to anything coming in contact with them. In this way,
mites further aid the spread of Trichoderma contamination. And, soon after an outbreak of
Trichoderma, it is not unusual to see a population explosion of mites.

Most notable are Trichoderma viride (a synonym of Trichoderma lignorum), an early appear-
ing mold with roughened spores and Trichoderma koningii, a smooth spored mold seen later in
the cropping cycle. Both are mushroom pathogens.

See Verticillium, a mold with similar symptoms when attacking fruitbodies.

See Color Photograph 22.

The Contaminants of Mushroom Culture/313

TRICHOTHECIUM

Class: Fungi Imperfecti
Order: Moniliales
Family: Moniliaceae
Common Name: Pink Mold.

Greek Root: From “trichos” meaning hairy
and “theke” meaning sac or capsule.

Habitat and Frequency of Occurrence:

For the most part, a saprophyte, rarely en-
countered in spawn making even though it is
one of the many microflora associated with
grain. Trichothecium is an occasional con-
taminant in agar culture and in poorly pre-
pared or immature composts.

Medium Through Which Contamina-
tion Is Spread: Primarily an airborne con-
taminant.

Figure 227 Drawing of conidia and sporu-
lating structure of Trichothecium.

Measures of Control: Air filtration and maintenance of good hygiene in the laboratory.

Macroscopic Appearance: Mycelium initially whitish; soon pinkish with spore production; and
typically slow growing on malt agars. Trichothecium is a powdery Penicillium type mold.

Microscopic Characteristics: Conidia measuring 12-18 x 4-10 microns; colorless to brightly
colored; two celled; pear shaped, ellipsoid or ovoid; borne in clusters with the basal cell being small-
er than the terminal one; and positioned at the apex of tall, thin, unbranched, but septate conidio-
phores. Spore bunches are attached to one another either in a chain-like fashion or in loose groups
but not lineally.

History, Use and/ or Medical Implications: One mold notable. Trichothecium roseum Link ex
Fr. secretes an antibiotic (trichothecin) that is toxic to bacteria, fungi and animals.

Comments. More frequently seen in the course of agar culture than on grain, this contaminant can
become a formidable problem if not detected early, and if large spore populations are permitted to
develop within the laboratory.

Also occurring on compost and occasionally on the casing soil, particularly where nitrogen

enriched compounds have not been converted into protein usable to the mushroom mycelium. By
itself it is not strongly inhibitory to mushroom mycelium, but thrives in habitats generally unsuited
for good mushroom growth. Adhering to good compost practices and following standard hygienic
procedures prevents this fungus from occurring.

See also Fusarium, a genus containing several pinkish colored contaminants and Geotrichum, a
genus known for the Red Lipstick molds.

The Contaminants of Mushroom Culture/315

VERTICILLIUM

Class: Fungi Imperfecti
Order: Moniliales
Family: Moniliaceae

Common Names: Dry Bubble; Brown
Spot; and Verticillium Disease.

Latin Root: From “verticillus” meaning
whorled or having branches on the same
plane, in reference to the shape of the conid-
iophore.

Habitat & Frequency of Occurrence: A

common parasite of the fruitbody.

Verticillium is promoted during cropping
under conditions of excessive humidity com-
bined with inadequate air circulation.

Verticillium grows within a broad Figure 228 Conidia and sporulating struc-
temperature range although warmer temper- ture of Verticillium.
atures (62 °F. and above) are preferred.

Singer (1961) reported an optimum of 72 °F. Verticillium abounds in soils and is introduced into
the growing environment via the materials composing the casing.

Medium Through Which Contamination Is Spread: Primarily transmitted from one infected
region to another by mushroom harvesters, flies and insects. Watering infected mushrooms further
spreads Verticillium spores.

Measures of Control: General hygiene maintenance; proper picking and cleaning practices; re-
moval or isolation of infected cultures; increasing air circulation; lowering of humidity; and elimina-
tion of flies and mites. If Verticillium is evident before a crop is harvested, carefully pick the infected
mushrooms, seal them in a plastic bag and leave the growing room with minimal contact with unaf-
fected areas. Verticillium spores are highly viscous and are best transmitted by motile hosts,
especially mites and other insects. Never water an infected bed until the diseased mushrooms have
been removed and the infected zones have been salted with alkaline buffer (baking soda, sodium
hypochlorite).

Macroscopic Appearance: Slightly infected mushrooms characterized by brown colored spots or

316/The Mushroom Cultivator

streaks on the basal or upper regions of the stem and on the caps of developing primordia. These
spots become grayish colored from spore production. Afflicted mushrooms often bend towards the
side that is infected. If the mushrooms do develop at all, they are typically tilted to one side or the
other. Verticillium attacks developing fruitbodies— the more severely infected are grossly mal-
formed, especially young primordia which are turned into sclerotia-like balls of amorphous whitish
mycelia. More mature but diseased mushrooms have a deformed pileus, sometimes with a “hair
lip”, and frequently with a downy grayish mycelium over the cap. The stem can be covered with a
downy mycelium and often vertically splits, roughly resembling a peeled banana. The cap becomes
disproportionately small relative to the fatter than normal stem. The overall texture of the mushroom
is dry and leathery.

When this mold attacks Psilocybe cubensis, there are several additional characters worthy of
note. Parasitized P. cubensis caps frequently become plane at an early stage. The stem becomes
swollen and hollow, narrowing radically towards the apex. Only in an extremely humid environment
does a downy mildew develop over the cap and stem surface. The “Verticillium spots” so com-
monly reported by growers of Agaricus, a white mushroom, are more accurately called “Verticillium
streaks” on P. cubensis, a mushroom with a brownish cap and a whitish stem.

Figure 229 Verticillium attacking Psilocybe cubensis.

The Contaminants of Mushroom Culture/317

Microscopic Characteristics: Conidia hyaline; unicellular; ovoid to ellipsoid; minute, measuring
1 -3 x 1-2 microns; borne singly or in small groups at the tips of narrow branches that whorl from a
central trunk at regular intervals. Conidiophores are slender and relatively tall.

History, Use and/or Medical implications: Apparently inocuous, no pathogenic species are
known.

Comments: Verticillium is the most common fungal disease parasitizing the mushroom crop and
the bane of both small and large scale growers. One misfortune of losing an early flush to
Verticillium disease is the increased probability of other diseases appearing. Split stems open the
mushroom up to attack by numerous insects and other pathogens. Not surprisingly, the sciarid fly is
a vector for the spread of Verticillium spores from parasitized mushrooms to healthy ones. It be-
comes clear that if conditions are right for Verticillium, the conditions are right for other molds. The
cultivator may soon have to deal with not one contaminant, but many.

Verticillium malthousei Ware is synonomous with Verticillium fungicola. Both are “brown
spot” fungi that envelope the mushroom with a fine grayish mycelium and cause brownish lesions
on their surfaces. Verticillium albo-atrum is another species in mushroom culture, although not as
frequently seen.

Verticillium is specifically a casing related contaminant that parasitizes the mushroom fruit-
body. Other molds that parasitize the mushroom are Dactylium and Trichoderma. They can be
separated microscopically. Dactylium spores are two celled and quite large (20 microns long) while
those of Trichoderma and Verticillium are single celled and much smaller (4x5 microns and 2-3
microns, respectively).

An easy method for the home cultivator to distinguish Verticillium infection from Trichoderma
is to plate out the suspect mold on malt agar media. If the mold is Trichoderma, forest green colo-
nies of mycelium will form. Other than green colonies of mycelium suggests the contaminant be
Dactylium or Verticillium.

Usually one sees Trichoderma blotch simultaneous to or after the occurrence of green mold
colonies on the casing layer. If there is no evidence of green mold on the casing layer and the mush-
rooms display these symptoms, then the mold is probably Verticillium or Dactylium. Dactylium
can be distinguished from Verticillium by its locus and manner of infection. Dacttyium is a grey,
aerial mold, fast growing and obvious on the casing. Verticillium is primarily evident on the fruit-
body and scarcely seen on the casing.

Steane (1 979) reported that Agaricus bitorquis seemed especially resistant to Verticillium dis-
ease whereas Agaricus brunnescens was more susceptible to it. Furthermore, he noted that farms
regularly suffering from this disease could greatly reduce the level of infection by intermittently
growing A. bitorquis between A. brunnescens crops.

A saprophyte and parasite causing “wilt disease” of many plants, particulary garden vegeta-
bles, Verticillium is abundant in most soils. Some Verticillium species are endoparasitic to nema-
todes— their spores germinate in the mouth tubes of the nematode with the resulting mycelia quick-
ly digesting the organism from within. Other pathogens that have similar symptoms to one or more
of the various stages of Verticillium are: Dactylium; Trichoderma; Mycogone; and Virus.

Pests of Mushroom Culture/319

M ushroom flies and midges are present in nature wherever fungi are found. Attracted by the
odor of decomposing manure and vegetable matter, as well as the smell of growing
mycelium, these insect pests zero in and lay their eggs on or near the mycelium and fruitbodies.
Under proper conditions these eggs hatch. But it is the larvae that do the extensive damage to the
mushroom plant, either by directly feeding on the mycelial cells or tunneling through the mush-
room fruitbody. Because of the concentration of attractive odors, a commercial mushroom farm is
always under siege by these pests. To insure insect free crops, certain measures are necessary. Un-
fortunately the bulk of these control measures involve insecticides, an approach not recommended
by the authors. The use of insecticides is not only costly and hazardous to human health, but also
represents a short term solution of a symptom rather than the solution of the problem itself. The
answer to disease and pest control in mushroom growing is strict hygiene for which there can be no
substitute.

Fly Control Measures

1 . Pasteurization periods and temperatures must be sufficient to kill all stages of insect
growth— 1 40 °F. for 2 hours in composts or other bulk substrates.

2. All Phase II, spawning, spawn running and cropping rooms must be airtight. Physically ex-
cluding insects from these areas is the most positive control one can exercize. Even the
smallest crack can serve as an entrance to the growing room. The spawn running rooms
should be the most secure with access to these areas restricted. All doors should be
weather-stripped and tight fitting. Positive pressure and air locks also help.

3. All tools and implements should be cleaned and disinfected before use on a new crop. A
commonly used disinfectant is a 2% chlorine solution.

4. Breeding areas must be prevented by removing from the premises all excess or spent sub-
strates, used grains, mushroom trimmings and other related by-products.

5. The growing room and all containers should be washed and disinfected between, crops.
Wood in particular harbors contaminants, including virus infected mushroom mycelium.
Treatment of wood with cuprinol or copper sulfate is common. Petroleum based products
should be avoided.

6. Fresh air intakes and exhaust vents must be screened with fine mosquito netting. Be sure,
there are no cracks around the filters and fan housing.

7. The room should be equipped with an insect monitor. The use of a monitor alerts the grow-
er to fly emergence from within the growing room or to fly entry from the outside. The
monitor can be as simple as a 1 2” x 1 2” plywood board to which a small black light (long
wave UV) is centrally mounted. On either side of the light sticky paper is attached. There
are also small pest lights commercially available. (See Resource section in the Appendix).

Pests of M ushroom Culture/321

Figure 231 Sciarid adult and its larva. (Adapted from P.R. VanderMeer; Penn. St.
Univ. Coop. Ext. Ser.)

Order: Diptera

Family: Lycoriidae (Fungus Gnats)

Genus/Species: Lycoriella solani, Lycoriella mali, Lycoriella auripila
Common Names: Sciarid Fly, Big Fly

Natural Flabitat: Predominantly saprophytes, living on wild mushrooms, rotting wood, leaf mold
and manure piles. Maturing mushrooms are frequently infested with sciarid larvae, the so-called
“worms” that commonly ruin choice wild edibles.

PHYSICAL CHARACTERISTICS

Mature Stage: Sciarids are small gnat-like flies characterized by two long segmented antennae,
large compound eyes, a black head and thorax and a yellow segmented abdomen. Females are
about 3 mm. long and can be distinguished by the swollen abdomen which ends in an ovipositor.
Males are about 2 mm. long and have a narrow abdomen ending in a distinct clasper.

Larval Stage: Larvae measure 6-12 mm. long with twelve abdominal sections and a distinct black
shiny head. The long creamy white body has a semi-transparent cuticle with a visibly darkened ali-
mentary canal. Larvae go through four development stages, or instars, before pupating.

Pupal Stage: Fully mature larvae spend two to three days spinning a cocoon of fine silky threads
and compost fragments. These threads are sometimes detected as slime trails left behind in the sub-

strate as the wandering larvae pupate. Once the cocoon is finished, the larva contracts into a pupal
stage, thus begining the transition to the adult stage. Pupae are 2-4 mm. long and change from
white to almost black.

Life Cycle: Developmental period in days

Temperature Egg Larva Pupa Adult

At 75 ° F 2 16 3 5-7

At 61 ° F. 7 23 8 (no data)

Sciarids thrive in the summer and fall with populations building to a peak in September and
October. Sciarids then die with the onset of cold outside temperatures.

Comments: The sciarid fly is responsible for considerable damage to commercial Agaricus crops.
Attracted by the smell of newly pasteurized compost, sciarids home in from miles away. A female
can lay between 150-170 eggs at a time. Eggs laid in the compost just after Phase II composting
hatch quickly into larvae during the spawn running period. These larvae then feed on the running
mycelium as well as compost, which is broken down into a foul smelling, soggy mass, totally unsuit-
able for spawn growth. Massive infestations can cause total crop failure.

At lower infestation levels, larvae migrate into the casing layer and then emerge just as the first
mushroom pins appear or as late as the first flush. These adults lay more eggs in the\cgj}ng, and the
newly hatched larvae attack both mycelia and mushrooms. Symptoms of this/a^|^3iclude:

1 . Dead pinheads.

2. Pins or mushrooms that are loosely connected to the casing due to severed mycelial con-
nections.

3. Brown or black spots on pinheads or on the stems of mushrooms.

4. “Salt shaker pins” perforated by larval tunnels.

5. Browning of the stem where cut.

Secondary damage to mushroom crops by sciarid flies comes from their role as carriers of
mites and diseases, including the pathogens Verticillium and Trichoderma. A single sciarid fly can
carry up to 20 mites!

Figure 232 Phorid fly and its larva. (Adapted from P.R. VanderMeer; Penn. St. Univ.
Coop. Ext. Ser.)

Order: Diptera
Family: Phoridae

Genus/Species: Megaselia nigra, Megaselia halterata
Common Names: Phorid Fly, Dung Fly

Natural Habitat: Commonly inhabiting manure piles and rank, decaying vegetation; feeding on
wild fungi and their mycelia. Phorid larvae are frequently seen tunneling through wild mushrooms.

PHYSICAL CHARACTERISTICS:

Mature Stage: Distinguishing features are a humped back, a rapid jerky run, a rounded third an-
tennal segment and a yellowish to reddish brown back. Adults measure 2-5 mm,, long. Females live
16 days and males live 10 days.

Larval Stage: Larvae are 6-10 mm. long, white and semi-transparent. The head is characterized
by a pair of “mouth hooks” with seven teeth. The segmented body tapers from the head to the pos-
terior end. Larvae pass through three instars.

Pupal Stage: Pupae are white at first then becoming pale yellow to brown. They can be distin-
guished by a pair of curved black respiratory horns.

Life Cycle: Developmental period in days

Temperature Eggs Larvae Pupa

75°F. 2 5 8

61 °F. 4 14 28

omments: Phorids can do extensive damage to the mushroom crop and are considered the prin-

324/Pests of Mushroom Culture

cipal mushroom pest in western Europe. Mated female phorids are drawn by the odor of mushroom
mycelium. This attraction increases during the spawn running period and peaks at full colonization.
Each female can lay up to 50 eggs which are placed in close proximity to the mycelium. In mature
mushroom crops, females lay eggs on the gills, in the casing, and adjacent to young pinheads.
Once hatched, the larvae feed on the mycelium, then tunnel into the mushrooms through the base
of the stem. Arising from these tunnels are secondary bacterial infections causing further damage
and brownish discolorations.

The fact that females will not lay eggs in total darkness gives the grower an effective method for
preventing Phorid infestation during spawn running.

Figure 233 Cecid fly and its mother larva. (Adapted from P.R. VanderMeer; Penn. St.
Univ. Coop. Ext. Ser.)

Order: Diptera
Family: Cecidomyiidae

Genus/Species: Heteropeza pigmaea, Mycophila speyeri.

Common Names: Cecids, Gall Midges

Natural Ffabitat: Commonly inhabiting decaying wood, rotting vegetation and manure piles or
wherever fungal mycelium occurs.

PHYSICAL CHARACTERISTICS:

Mature Stage: Adult cecids measure less than 1 mm. long making them almost invisible to the
naked eye. H. pigmaea are orange with a long segmented abdomen and segmented antennae.
Wing venation or structure is noticably absent except close to the thorax.

Larval Stage: Newly born larvae are 1 mm. long and 2-3 mm. when mature. H. pigmaea are
white to cream; M. speyeri are bright orange. Larval movement is facilitated by free water,
whereas in dry conditions this movement is by flexion, jumping as far as 2 cm. Larvae are photo-
kinetic (moving to light) and can reproduce through paedogenesis, a process whereby mother lar-
vae give birth to daughter larvae. Under optimal conditions mother larvae can produce 14-20
daughter larvae in six days. Thus, in a short period of time a population explosion can occur.

Pupal Stage: H. pygmaea larvae molt to a rigid “hemi-pupa” wilhin which new daughter larva
evolve. Conditions favorable to larval growth lead to a “resting mother larvae” stage which can

326/Pests of Mushroom Culture

remain alive up to 1 8 months. M. speyeri has neither of these particular attributes although it also
performs paedogenesis. Larvae of both species can change to “imago” larvae, form only one in-
star, then molt to free pupae, emerging as adults five days later.

Life Cyc le: Developmental Period in Days

Egg Mother Larva Daughter Larva

2 E/6 (2) E/6

Comments: Cecid larvae pierce or tear growing hyphae, sucking out the contents. The main
loss suffered by commercial growers is contamination of the mushrooms by larvae. H. pygmaea
can also carry a bacterium which produces longitudinal brown stripes on the stem. In the infected
mushrooms, tiny black droplets of fluid form on the gills, which then become spotted or turn
black.

Figure 234 Wing venation of mushroom flies: clockwise from top right, Leptocera,
Sciarid, Cecid and Phorid.

Order: Dipt era

Family: Sphaeroceridae (Borboridae)

Genus/Species: Leptocera heteroneura

Natural Habitat: Associated with manure, compost piles and decaying organic matter.

PHYSICAL CHARACTERISTICS:

Mature Stage: Leptocera has large red compound eyes and with a yellow and black striped ab-
domen. Leptocera flies are very similar to phorid flies but are smaller and have a distinctive wing
venation. They somewhat resemble the common fruit fly.

Larval Stage: Larvae have a blunt posterior end tapering to a slender head which is equipped
with mouth hooks. Leptocera larvae are very similar to house fly maggots in appearance.

Pupal Stage: Pupae are golden brown and barrel shaped.

Life Cycle: Developmental Period in Days .

Egg Larva Pupa

3 14-28 10-14

Comments: The Leptocera fly acts as a vector for disease organisms and is frequently associated
with bacterial infections. It is a known carrier of mites.

328/Pests of Mushroom Culture

Mites are very small spider-like insects that live and breed in decomposing vegetable matter,
feeding on molds present therein. Optimum breeding environments are moist and warm, giving
rise to a rapid succession of generations and exponential growth. Under adverse conditions cer-
tain mites have the ability to change into an intermediate stage called a “hypopus”. The hypopae
have flattened bodies, short stubby legs and a sucker plate with which they attach to moving ob-
jects. These attributes facilitate dispersal. An excellent survival mechanism, it is the hypopae that
are commonly carried by flies. A typical life cycle for mites in days is:

Temperature

Eggs

Larvae

Protonymph

Tritonymph

Total

75 °F.

60 °F.

1 1

Mites are known to eat mushrooms and their mycelia. Additionally they devalue the crop and
crawl onto pickers, causing temporary discomfort. Their presence is an indication of unsatisfac-
tory substrate preparation and insufficient pasteurization times and/or temperatures.

Figure 235 Straw mites.

Pests of Mushroom Culture/329

Order: Arcana
Family: Tyrogelyphidae

Genus/Species: Tyrophagus putrescentiae, Caloglyphus mycophagus
Common Names: Straw or Hay Mites

Discussion: Straw mites have soft translucent pinkish or yellowish bodies punctuated by long
flexible hairs. One female is capable of producing 500 eggs in a lifetime. Commonly found in hay
or straw piles, these saprophytic mites are endemic to foul compost. They feed on molds and
bacterial contaminants of the mushroom crop and also eat mycelium and mushrooms, making
small irregular pits in the stem and cap. These pits can later become infected by bacteria.

Order: Arcana
Family: Eupodidae

Genus/Species: Linopodes antennaepes
Common Name: Long Legged Mushroom Mite.

Discussion: This mite is easily recognized by its long front legs which are twice the length of the
light, yellowish brown body. It is not believed to be directly injurious to the mushroom crop and in
fact is a predator on other mite species.

Order: Arcana
Family: Tarsonemidae

Genus/Species: Tarsonemus myceliophagus
Common Name: The Mushroom Loving Mite.

Discussion: Tarsonemus mites are very small, 180-190 microns long, with pale brown, shin-
ing, oval bodies. They occasionally swarm in masses on mushroom caps but otherwise are rarely
seen except by microscopic examination. Females produce an average of 22 eggs in a lifetime of
2-8 weeks. These mites cause a bright reddish-brown discoloration at the base of the mushroom
stem and may cut the stem’s mycelial connections. Known to survive normal compost pasteuriza-
tion temperatures, they can carry a virus disease to Agaricus brunnescens.

Order: Arcana
Family: Pyemotidae
Genus/Species: Pygmephorus sp.

Common Names: Red Pepper Mites; Pygmy Mites.

Discussion: Pepper mites are small (250 microns long) with yellowish brown, wedge-shaped
bodies, crossed by a central whitish band. Red pepper mites are often seen as a swarming jostling
mass, on mushroom caps or the surface of the casing. These mites are commonly associated with
Penicillium and Trichoderma molds, upon which they feed.

330/Pests of Mushroom Culture

WATER

SAMPLE

■FUNNEL

Figure 236 Light micrograph of Red
Pepper Mite. Note that darkened shapes
by front leg are Panaeolus subbalteatus
spores. See also Figure 230.

Figure 237 Nematode testing apparat-
us. Sample is wrapped in gauze and
submerged in a water filled funnel. After
twenty-four hours, a small amount of
water is drawn off and examined with a
magnifying lens or dissecting scope.

Pests of Mushroom Culture/331

Nematodes or eelworms are microscopic roundworms which live in soil, decomposing
organic matter, fresh or salt water, or on living host plants, fungi, insects and animals. Nematodes
can survive up to six weeks without food and are unaffected by freezing. With eight billion
nematodes in each acre of soil, they are one of the most numerous creatures on earth.

Water is essential for locomotion and breeding. Swimming in an eel-like fashion and because
of their minute size, nematodes can live in the thinnest films of water. With sufficient water, nema-
todes rise to the surface of their environment. In moist casing, large numbers of nematodes are
visible as a shimmering veneer on the casing surface. This behavior is called “winking” and is
caused by the nematodes standing on their tails and waving their bodies in the air. Considered to
be an adaptation for dispersal, the winking nematode adheres by means of a sticky outer skin to
whatever they come in contact with, be it a fly, mite, human hand or clothing. This same outer
skin protects the nematode from adverse conditions.

If dried slowly, nematodes can change to a “cryptobiotic” or “cyst” state, thereby preserved
for years until reactivated by water. In this cyst state, nematodes are also able to persist in high
temperatures that would otherwise be lethal.

Parthenogenesis, the ability for females to breed asexually without males, is common among
nematodes and leads to very rapid population expansion. By this means, a single nematode can
breed millions of descendants within a few weeks. Nematodes can also reproduce sexually, but
not as rapidly.

Nematodes present in mushroom culture can be classed into two basic types according to
their feeding habits: saprophagous and mycophagous.

Saprophagous Nemalotodes

Genus/Species: Rhabditus spp.

Saprophagous eelworms are characterized by a tube-like mouth through which they suck nu-
trient particles suspended in water. These nutrients are comprised of organic matter and its
accompanying microorganisms, particularly bacteria. Because bacteria occur in large numbers in
both mushroom compost and casing soil, these materials provide excellent breeding grounds for
saprophagous eelworms.

In bulk substrates such as compost or plain straw, nematodes can be found in great numbers.
The high temperatures of Phase I conditioning would normally destroy them if it were not for the
fact they migrate to the cooler outer shell of the compost pile. Phase II can eliminate nematodes
but only if the entire compost is subjected to pasteurization temperatures. In a properly prepared
and thoroughly pasteurized substrate, the mushroom mycelium consumes all free water and then
feeds on the bacterial population. This creates a “bacteriostatic environment”, which effectively
limits nematode growth capabilties. In an uneven substrate with overly wet and dry areas,
however, the nematode’s ability to breed increases. Wet areas are particularly suitable for

332 / Pe sts of Mushroom Culture

Figure 238 Mycophagous eelworm (top) and Saprophagous eelworm. Note stylet in
mouth tube of former.

eelworms to breed and feed. And, as their population increases, the build-up of waste material
from metabolic excretions soon fouls the substrate, rendering it unsuitable for mycelial growth.
These excretions result in similar damage to infested casing soils.

Although saprophagous eelworms are not primary pathogens, their presence indicates im-
proper hygiene or imbalanced growing conditions. For this reason, control measures focus on
prevention rather than treatment. In fact, there are no practical means to treat infested areas that
would not likewise harm the mushroom mycelium.

Mycophagous Nematodes

Genus/Species: Ditylenchus myceliophagus; Aphelenchiodes composticola

Mycophagous eelworms feed directly on mushrooms. They are characterized by a mouth
stylet or needle with which these eelworms puncture hyphae, inject digestive juices and then suck
out the cellular contents. The damaged cell, drained of its cytoplasm, soon dies. Feeding continu-
ally and moving from cell to cell, mycophagous eelworms can soon destroy whole mycelial net-
works. In infected substrates, the fine mycelial growth disappears, leaving only the coarse strands
which give the appearance of stringy growth. Eventually the substrate becomes soggy and foul
smelling, a condition further promoted by the build-up of anaerobic bacteria. Often the nematode
trapping fungi, Arthrobotrys spp. develop in association with them. It is visible as a fine grayish
mold-like growth. Although the presence of this mold is a useful indicator of nematode infestation, it
is not a true control for these organisms.

Mycophages differ from saprophages in their slower non-parthogenetic reproduction and their
lack of the “winking” behavior mentioned earlier. Both Mycophagus species can reproduce
30-100 fold in about two weeks at 70-75 °F.

Mushroom Genetics/333

■■

Figure 239 Giil face of Psilocybe cyanescens populated with fertile spore-bearing
basidia and sterile cells called pleurocystidia.

334/Mush room Genetics

T his chapter discusses what genes are and what they do. It addresses the relationship between an
individual’s set of genes and the characteristics of that individual. The implications of genetics
for the grower or breeder of mushroom strains are examined and an improved, easy technique for
generating cultures from spore prints will be presented.

What Are Genes?

Genes contain specific sequences of nucleotides, the nitrogen-based building blocks of the
DNA molecule. These sequences specify the order of nucleotides in messenger RNA molecules,
which in turn determine the sequence of the amino acids in a protein chain. For the purposes of this
discussion, genes may be regarded as indivisible units, although in fact, they can on rare occasions
be split or altered. A mutation is the permanent alteration of a gene caused by some outside force
(chemicals, radiation, mistakes by the DNA copying mechanism of the cell, etc.). In discussions of
genetics, a gene is often referred to as a genetic locus, emphasizing the fact that genes are regions
of a DNA molecule. Within a population of a species, there are many differing copies of each gene.
Each copy is referred to as an allele of that gene.

What Do Genes Do?

Genes are the blueprints of life. They specify the structure of RNA and protein molecules; these
molecules create all the other compounds and structures which make up a living organism. An indi-
vidual organism is an emergent property of its genes in that not only is it the result of gene products,
but also the interactions of gene products. The expression and interaction of genes, that is the char-
acteristics of an individual, are known collectively as the phenotype of that individual, whereas the
sets of genes which produce the phenotype is known as the genotype.

The Advantage of Multiple Copies of Genes

Many genes are present in the genotype in several copies, and these copies are often different
from one another. This is because the protein specified by any one gene copy has unique physical
and chemical properties of its own. It functions most efficiently at a certain temperature, pH and salt
concentration. If an important protein is represented in several different gene versions, a broad band
rather than a narrow range of temperatures and chemical conditions will be optimal.

Chromosomes

Chromosomes are collections of genes. They are long DNA molecules, each of which contain
several thousand genes. For this discussion, the genes are best visualized as beads on a string, so
that the string can be cut at any point between the beads, and can be rejoined at the place of the cu<
or to any cut “string portion” or the chromosome.

Chromosomes are very small. With special stains and high powered microscopes, the larger
ones can be seen. Unfortunately, the chromosomes of fungi are extremely small, and the number of
chromosomes, something characteristic of each species, has never been determined for most fungi

Mushroom Genetics/335

There is a complete set of chromosomes in every cell of every organism. This means that every
time a cell divides, a complete copy must be made of every chromosome, and hence of every gene
in the organism’s genome. The cellular copying process is very nearly perfect, with errors being
made at about the rate of one per million genes. That is , to find a random mutation of a particular
gene, you would have to look at a million cells. Factors which produce mutations will, of course, in-
crease this rate. These copy errors are the source of background mutations, which are always ap-
pearing in every organism.

Mitosis

Mitosis is the normal process of chromosome duplication which takes place every time that a
tell divides. In it, all the chromosomes are duplicated, and in the early stages of the process, the
copies stick together. All of the duplicated chromosomes line up in the middle of the cell, and one
of the two copies of each is pulled to either end of the cell, resulting in two complete sets of chromo-
somes.

Meiosis

Meiosis is the unique series of events which takes place when a cell is involved in sexual repro-
duction. In meiosis, the chromosomes are copied just as in mitosis, but the genes are shuffled in a
process called recombination. Sexually reproducing organisms have two sets of chromosomes,
one from each parent. In meiosis, these two sets line up side by side, and reciprocal exchanges of
sections of chromosomes take place. That is, a section of the maternal copy of a chromosome is
transferred to the paternal copy, and a section of the paternal copy is simultaneously transferred to
the maternal one. This happens to all of the chromosomes, usually once per chromosome, but
sometimes more than once.

After these reciprocal exchanges take place, two successive cell divisions occur, resulting in
four cells, each with ONE copy of each chromosome. None of these cells are identical to any of the
others. They each have unique sets of genes. These cells are known as gametes, and are basidio-
spores in a mushroom, ascospores in a cup fungus or a yeast and sperm or egg cells in an animal.

It is this act of recombination of genes within the genome and the combination of genomes
from two parents which is the genius of sexual reproduction. By this mechanism, variety is constant-
ly introduced into the population of a species. A bacterium, which can reproduce very rapidly by mi-
tosis, can generate vast numbers of bacteria in a very short time, but all of the offspring are iden-
tical. The importance of this difference cannot be overstated. If conditions fall below optimal or into
the lethal range for the parent bacterium, all of the progeny soon die or are equally affected (unless,
of course, there has been a favorable random mutation). Chance favorable mutations are, in fact,
the major means of evolution available to bacteria.

Sexual reproduction, on the other hand, constantly spins off variation. Some of the progeny are
substandard and do not survive, or do poorly, most are average and some are clearly superior, flour-
ishing and leaving behind a greater number of offspring than the other groups. In this way, the pop-
ulation is enriched in gene combinations which are better adapted to the environment.

336/Mushroom Genetics

The two aforementioned modes of reproduction lead to three primary reproductive strategies.
These are the primary use of asexual reproduction, the primary use of sexual reproduction and the
sequential or seasonal use of both methods of reproduction.

1 . Asexual (mitotic) reproduction allows an organism to produce large numbers of offspring in
a very short period of time. This makes possible the rapid exploitation of any ecological niche which
becomes available. This strategy is used by bacteria, yeasts, many molds (Fungi Imperfecti) and a
surprising number of plants.

2. Sexual reproduction is not as rapid, since meiosis, gamete production and fusion and zy-
gote growth are relatively slow processes. The progeny, however, have built-in variation and are ca-
pable of exploiting a wider assortment of niches than the parents. This strategy is used by larger or-
ganisms which tend to live for a longer time than those which are primarily asexual. Examples of or-
ganisms using this strategy are polypores, most plants and all large animals.

3. Combining sexual and asexual reproductions in different portions of the life cycle results in
a highly effective strategy. This method is utilized by most lower plants and most fungi. In this strate-
gy, when a suitable niche is found, asexual reproduction allows it to be rapidly filled and exploited.
When that niche has been populated and nutrients become scarce, sexual reproduction is triggered.
As well as releasing a number of varied progeny to the environment, sexually produced spores are
usually more resistant to the harsh environmental conditions than mitotically produced spores.
Often they are specifically adapted to lasting through winter or through a period of dryness, condi-
tions not conducive to the growth of fungi.

Asexual Reproduction in the Fungi

Asexual reproduction in the fungi takes many forms, including buds, conidia, sporangiospores
and fragmentation products.

Yeasts reproduce by budding, which is the constant growth of new cells from the surface of a
mother cell. The new cells literally “blow out” of the mother cell wall like a balloon.

Conidia are mitotic spores which are continuously produced within or upon special structures
called conidiogenous cells. Examples of conidial fungi are represented by Penicillium and
Aspergillus molds, the fungi which attack spoiled foodstuffs, the downy and powdery mildew which
attack garden plants, and the hundreds of genera which are involved in the breaddown and re-
cycling of debris and litter in nature.

Sporangiospores (spores formed in batches within saclike structures called sporangia) are
found in the water molds and the Zygomycetes. Rhizopus, which is often seen on breads and straw-
berries, reproduces in this manner.

A common mode of asexual reproduction is for portions of vegetative mycelium to thicken and
form heavy walls and septae. These reinforced hyphal fragments then break apart and are distri-
buted by natural processes. These vegetative propagules are called by many names, including ar-

Mushroom Genetics/337

throspores, chlamydospores, gemmae and others.

Sexual Reproduction in Mushrooms

While mushrooms reproduce sexually, they have no sexes. All that the term sexual reproduc-
tion means is that two sets of genetic information are carried, and that the genes in those sets are
shuffled randomly before one set is provided to each gamete. Two gametes must come together
and fuse to form the next fertile generation.

In animals and plants, the notion of sexes is realistic, because there are two kinds of gametes,
an egg and a sperm. In mushrooms, all the gametes are physically identical; they are the basidio-
spores. Because of meiosis, however, there are genetic differences between them.

One of the genetic characters sorted out during meiosis is the mating type. The mating type is
a character which prevents a spore or monokaryotic hypha carrying a particular allele from fusing
sexually with any spore or hypha carrying the same allele, no matter how different the genomes are
at all other loci. It takes the presence of different alleles at the mating type locus for sexual reproduc-
tion to occur. In any one species, there may be any number of alleles within the population. In gen-
eral, any one of them is compatible with all of the others, the only prohibition being against fusion
with the identical mating type.

If a species of mushroom has only one locus controlling mating type, with varying numbers of
alleles for that locus, that species has what is known as an unifactorial, heterothallic mating sys-
tem. In such a system, the only physiological requirement for mating to take place is that two differ-
ing alleles of the mating type locus be present. Since two alleles must be present in a sexually ma-
ture mushroom, and each spore only gets one, any randon spore is compatible with half of its sib-
lings. Since there are a large number of alleles for the mating type locus in the population at large,
any random spore has a higher probability of being compatible with a spore of another strain. Thus
this system increases the percentage of outcrossing by members of the species using it.

The majority of mushrooms, however, are heterothallic and bifactorial, a system known as
tetrapolar. In this system, there are two separate and distinct mating type loci, each of which must
have differing alleles present to form a dikaryotic colony. This system produces four distinct types of
spores on each basidium, and any random spore from a single strain is fertile with only one fourth of
its siblings. This is a strong form of incest taboo, and makes it four times as likely that any naturally
formed dikaryon will be from non-related spores. Unfortunately, the two types of spores which are
not totally identical or non-identical can form dikaryotic colonies which look like fertile ones. These
products of illegitimate matings, though, are incapable of making fruitbodies or basidiospores.

There are strains and species in which the mating type system has broken down. These are
known as homothallic fungi, and they are fully capable of mating with themselves. In fact, a single
spore of a homothallic fungus is usually capable of making a ferile dikaryotic colony. A fair number
of spores, however, due to the effects of recombination, will be incapable of forming fertile colonies
unless they mate with another strain. This is a system often found in fungi which live in marginal
habitats; usually there is a time lag before a monokaryotic colony dikaryotizes itself.

338 /Mushroom Genetics

There are two types of homothallism in mushrooms: primary and secondary. Primary
homothallism is the case described above, where the majority of spores, while initially forming
monokaryotic colonies, will eventually become dikaryotic and fruit normally. Secondary homothal-
lism is the case where each spore receives one nucleus of each mating type, generating a dikaryotic
colony from the moment of spore germination. Agaricus brunnescens is the best known example
of this type of fungus, while another commonly cultivated mushroom, Volvariella volvacea, has a
primary homothallic mating system.

The single most important implication of the genetics that has been described thus far is the oc-
currence of illegitimate matings. In a tetrapolar fungus, only one fourth of the spores from any one
mushroom are fully compatible with any random spore from that same strain. This mechanism ex-
ists to encourage outcrossing. When a cultivator is trying to produce a strain from a spore print, es-
tablishing a fruiting strain can be frustrating. This is because monkaryotic hyphae with common A
factors or with common B factors can fuse and form dikaryons, and these dikaryons can even
make convincing looking clamp connections. (See Figs. 10 and 182). These colonies, however,
are incapable of fruiting. It becomes obvious at this point that two thirds of the random dikaryons
formed will be of the illegitimate type. This implies that a large number of dikaryotic cultures must
be isolated and tested for fruiting ability. Another, but less precise way around this problem is to
inoculate with a large number of spores and take a tissue culture of the first mushroom that appears
in the culture. This procedure is the one usually listed in books on mushroom cultivation because it
is simple, but the strains produced in this manner still must be tested thoroughly.

The phenomenon of sectoring is the production of wedge shaped areas of differing physical
or growth characteristics by a colony of mycelium. There are two types of sectoring, one found in
young cultures, and one in old ones.

In young multispore cultures, several different strains are all growing together at the same time.
Some are the products of legitimate matings, some of illegitimate ones. These strains all have differ-
ing characteristics. Some of these strains grow faster than others, some are rhizomorphic and some
are fluffy in appearance. Some fruit well, some poorly. Some produce clumps of many tiny mush-
rooms, some produce a few large ones. They each have a unigue set of preferred culture condi
tions.

Fortunately, the different strains formed from multispore germinations tend to sort themselves
out. As the colony grows, strains segregate into sectors of different appearances and growth rates.
The repeated separation and propagation of individual sectors, until a colony is obtained which no
longer produces new ones is one way of isolating a pure strain. Several strains may be isolated from
the same original petri plate in this way.

As pure cultures grow old and become senescent, they produce ever greater quantities of sec-
tors due to the accumulation of random mutations. Repeated subculturing of the culture gives accu
mulated mutations a chance to express themselves. A strain which has reached this condition is no
longer pure, and should not be used for cultivation.

Mushroom Genetics/339

Culture Trials

When a number of strains have been generated from a sporeprint, they are different because of
recombination in the basidium. Some of the strains MAY be identical to the parent strain, but that
must demonstrated by some testing procedure. As in any screening operation, the more strains
used, the better the chance of a good result. In fact, professional mushroom breeders often do trials
with thousands of strains at a time. This kind of work, however, takes large and expensive facilities,
and is unnecessary if the purpose is simply to find a strain which fruits well under a certain set of
conditions. A strain which fruits well in test batches under uniform conditions has a high likelihood
of doing well in larges batches when the same conditions of temperature, humidity and aeration are
maintained. How many strains need to be tested? If the mushroom being worked with is tetrapolar,
only one third of the dikaryotic colonies picked out will be capable of fruiting at all. In order to make
trials of ten fruiting strains, begin with at least thirty dikaryotic strains.

Many mushrooms, especially the wood-rotters, fruit on enriched agar media in a petri plate if
given proper temperatures and some light. If the mushroom being tested is one of these, the selec-
tion of fruiting strains is simple. A mushroom requiring a special substrate or additive to fruit should
be provided with the smallest amount allowable. For example, Agaricus brunnescens can be fruited
on 50 grams of sterilized grain in a pint jar, if it is cased with soil containing certain bacteria. The

Figure 240 Two spored basidium of a Copelandian Panaeolus.

340/Mushroom Genetics ^ __ , ...-bee

smallest possible amount of substrate allows the rapid determination of fruiting strains.

Once ten to fifteen fruiting strains are in hand, they should be tested in a small scale version of
the ultimate culture method. This step allows the strain best adapted to culture conditions to be se-
lected. All strains should be tested at least in duplicates; five replicates per strain are preferable.

If the ultimate cultivation method involveI||>eds of compost, the tests can be made with small
boxes filled with compost, but the boxesj i£»|j}>e filled to the same depth as the beds will be in the
full scale project. If the fungus is fruitedWw$fex. Flammulina velutipes), a few jars can be inocu-
lated with each strain. Good records musfSe iVJjfjt for comparing the fruiting potential of each strain .

In small scale trials such as these, often several strains look good. In this case, the only way to
find the best one is to make full scale trials, with one third or one fourth of the jars or beds inoculated
with each of the strains being tested. Once again, good record keeping practices should soon show
the differences between the most and least productive strains.

If the mushroom under consideration for cultivation takes a long time to establish its fruiting
cycle (ex. Lentinus edodes ), it is best to simply purchase a culture from a spawn lab or to take tissue
cultures from commercially grown mushrooms.

Spore Dilution Technique

A simple technique can be used to physically separate spores so that individual dikaryotic (or
even monokaryotic) cultures can be isolated in one step. The necessary equipment includes a bac-
terial (small) inoculating loop, several screw-cap vials of 20-30 ml. capacity, a flame and several
sterile pipettes or small syringes.

To utilize this method, first fill each of the vials with 9 ml. of distilled water, place the caps on
loosely and sterilize them. After they have cooled, the caps should be firmly screwed down. The in-
oculation loop is then flamed, cooled and gently rubbed on the spore print, being careful not to get
a large mass of spores. The loop is dipped and twirled in one of the vials, which is then recapped
and shaken vigorously. One milliliter of the fluid is then transferred to another vial, which is re-
capped and shaken, generating a dilute spore suspension. This suspension may be further diluted in
the same manner. In this way, the cultivator has generated three suspensions of spores, one of high
spore density, one 1 /1 0th as concentrated and one with 1 / 1 00th or 1 % of the original concentra-
tion. Now spread 1/1 0th of a mililiter of each suspension on a separate media filled petri plate (or
better yet, use several plates for each dilution). The original strength suspension in all likelihood will
produce a dense lawn of cultures, which will be difficult to separate. This is the same condition as is
produced with normal spore spreading methods. The less dense suspensions, however, should pro-
duce many fewer colonies, usually in the range of 20-50 per plate for the 1 : 1 0 dilution and 2-5 per
plate for the 1 :1 00 dilution.

Look carefully at the plates having only a few colonies. The slower growing monokaryons can
be discerned from the faster growing dikaryons. Pick about 25 of the dikaryons to test for fruiting
ability and reaction to culture conditions.

If desired, the monokaryotic cultures can be picked out for a breeding program. This is espe

Mushroom Genetics/341

dally valuable if there are spores from several strains available. When spores are simply spread onto
a plate, they adhere to one another, so attempts to simply streak spores of two strains on a plate usu-
ally do not yield hybrids.

(The authors gratefully acknowledge Michael McCaw for the contribution of this chapter on
genetics).

342 /Mushroom Genetics

Appendix I: Medicinal Properties/345

M ushrooms have long been esteemed for their medicinal properties, especially by Far Eastern
cultures, while western cultures have largely been oblivious to the beneficial properties of
mushrooms. For centuries, the Japanese have hailed the shiitake mushroom ( Lentinus edodes) as
an elixir of life, a cure-all, revitalizing both body and soul, a cure for cancer, impotency, senility and
a host of other ailments. Mazatec shamans of southern Mexico have used Psilocybe mushrooms in
their divination and healing ceremonies, extolling them for their life-giving properties and calling
them “Mushrooms of Superior Reason” for the heightened mental state they induce. Even the very
term “agaric,” still used to describe all mushrooms with gills, comes from the name of a pre-
Scythian people, the Agari, who were skilled in the use of medicinal plants, of which mushrooms
were one.

Not until the late 1920’s, when Dr. Alexander Fleming published a note in a microbiological
journal, did fungi draw the scrutiny of scientists looking for new sources of antibiotics. He observed,
quite by accident, the deterrent effect a Penicillium mold had on a bacterial contaminant (a
Staphylococcus species). Years later, fellower researchers pursued his suggestion that antibiotics
were being produced by this mold, which shortly led to the discovery of penicillin. Forthwith, molds
of all types were examined by W.Ff. Wilkins (and others) from 1 945 to 1 954 who systematically
tested one hundred species at a time for antibiotic effects against bacteria and bacteria-carrying
viruses. Eventually, Wilkins turned his attention to the fleshy fungi and interest within the scientific
community grew.

Claims of healing properties in mushrooms have been primarily promoted, until recently, by
the commercial mushroom industry and others with vested interests. It appears, however, much of
the medicinal claims attributed to mushrooms are not myth, but founded in some truth. Within the
last ten years, numerous studies demonstrating the anti-cancer and interferon stimulating properties
of Lentinus edodes have been published. Individuals can significantly reduce serum chloresterol
levels by eating these mushrooms for as short a period as a week (Suzuki and Ohshima, 1 974). In
another study (Hamuro et al. , 1974), the antitumor influence of hot water extracts of Lentinus
edodes was demonstrated in mice implanted with sarcoma-180 and other cancers, resulting in a
80% remission from treatment lasting only ten days, and a 1 00% prevention of growth if the mice
were injected prior to implantation. The causal compound is appropriately named ientinan, a anti-
tumor polysaccharide. Extracts from shiitake spores and the isolation of “mushroom RNA” from
them have proved effective against influenza (Suzuki et al., 1974). Similar antitumor, immuno-
potentiator and interferon stimulating polysaccharides have been found in Boletus edulis, Calvatia
gigantea, Coriolus veriscolor, Flammulina velutipes, Ganoderma applanatum, Ganoderma

lucidum (the classic “Reishi Mushroom”), Phelinus linteus, Armillaria ponderosa (Tricholoma
matsutake) and Pholiota nameko. (See Yamamura and Cochran, 1974).

In the treatment of other diseases, Cochran and Lucas (1959) reported Panaeolus sub-
balteatus, a mushroom producing psilocybin and psilocin, provided significant protection from
polio virus in mice as did several other edible and inedible mushroom species. Psilocybian mush-
rooms might be of further usefulness in improving eye sight, hearing, circulation and in activating
the self-healing processes within the human body.

With the current emphasis on prevention and natural cures for human diseases, mushrooms
are proving to be a convenient, inexpensive and an effective method of sustaining health. Health
conscious individuals beginning a daily regimen of eating shiitake, for instance, have been shown to
be less suceptible to virus-induced diseases than those abstaining. Until these studies progress and
are tested more extensively on human populations, hopes should not be unduly raised for mush-
rooms might be of further usefulness in improving eye sight, hearing, circulation and in activating
the self-healing processes within the human body,

Appendix II: Laminar Flow Systems/347

S uspended in the air is an invisible cloud of contaminants. These airborne spores are the primary
source of contamination during agar and grain culture, and they are the major force defeating
beginning cultivators. To control contamination, the cultivator must start with a sterile laboratory.
Without pure culture spawn, the prospect for a good crop is slight, no matter how refined one’s
other techniques.

Creating an absolutely sterile environment, free of all airborne particulates, is extremely difficult,
if not impossible. “Nearly sterile” environments are more easily constructed and are quite suitable
for the purposes of the mushroom cultivator.

Chemical cleaners like detergents and disinfectants have traditionally been used for this pur-
pose. Unfortunately, the frequent use of these cleaners to maintain hygiene in the laboratory pose
some risk to the handler. Ultraviolet lights are likewise dangerous and are difficult to position in a
room so that no shadows are cast. By far the least harmful and most effective method is the use of
high efficiency filters that screen out airborne particulates when air is pushed through them. These
filters are the basis of laminar flow systems. An understanding of the composition of unfiltered air
helps put into perspective the problem for which laminar flow systems are designed. The air, the fil-
ter, the fan and the laminar flow system will be discussed in that order.

The Air

Air is composed of many suspended and falling particles. A sample of air holds soot or smoke,
silica, clay, decayed animal and vegetable matter, and many, many spores. Some are only a fraction
of a micron in diameter while other are hundreds of times larger. These particles continously rain
down on the earth’s surface. In light impact zones isolated from industrial centers, twenty tons per
square mile per month fall from the sky (ASHRAE, 1 978). Industrial areas have a fall-out that is ten
times greater. So-called “clean country air” contains, on the average, one million particles (greater
than .3 microns) per cubic foot. But in a room where a cigarette is being smoked, more than one
hundred million particles are suspended in the same air space. A sterile laboratory, on the other
hand, has less than one hundred particles per cubic foot of air!

Most of the spores contaminating mushroom cultures are between .5 and 20 microns in diam-
eter. Generally, particles greater than 10 microns fall out of the air because of their weight. The
smallest particles in this group are the airborne spore-forming bacteria which originate from soils.
The smallest endospore forming bacteria are around .4 microns in diameter. Viruses which meas-
ure even smaller, sometimes a mere .05 of a micron in size, are usually attached to larger particles
such as fungal spores. This broad assortment of airborne debris poses the greatest danger to mush-
room culture.

Figure 239 A standard design of a laminar flow cabinet for tissue culturists.

The Filter

Two types of high efficiency filters are available today. One is an electrostatic filter which will
screen out spores down to 5 microns or less. These filters operate on a charged particle principle
where, by a variety of means, airborne particles are passed through an ionizing field and then be-
tween two oppositely charged electrical plates. Charged particles are drawn to the grounded plate
by the force of the electric field. Because an agglomeration of particles is likely to blow off the retain-
ing plate, they are often coated with a special oil. The advantages of electrostatic filters are that they
have little resistance (a low pressure drop) and that they are reusable. But they have several disad-
vantages. One disadvantage is that they do not screen out the particles of 1 micron or less with a
99 + % efficiency in high velocity airstreams. Hence, as air velocity increases, their efficiency de-
creases. Many electrostatic filters have, as a result, a sliding scale of efficiencies based on air speed.
Another problem associated with electrostatic filters is that particles not caught in the filter are still
partially charged and stick to the walls of a room, discoloring them. Also, toxic ozone may be gener-
ated by the constant arcing in the electrostatic field.

The basic element in an air filter is the media, particularly the dry extended surface kind that is

Figure 240 A commercially available laminar flow
hood.

rated to .3 or .1 microns. Extended surface filters are commonly known as HEPA (High Efficiency
Particulate Air) filters. First used commercially in 1961, these filters are honeycombed with fine
sheets of microporous material that can screen out particulates down to less than one third of a
micron size with a rated 99.99% efficiency. All spores of plants, fungi and most bacteria are thereby
trapped within the folds of the filter.

The collection media in this type of filter can be composed of various materials including hair,
spun glass, wool, paper and asbestos. (In the past, asbestos has been used in the manufacturing of
all types of filters. Since asbestos is cancer causing, be sure to specify a non-asbestos fiber). The ex-
tended surface media filter consists of folds of material woven back and forth. Corrogated aluminum
or paper separators are inserted perpendicularly to the filter face and separate the folds to help direct
airflow in an even, parallel fashion.

The airstream hits the filter material at a perpendicular angle and is forced to pass through the
many weaves of the filter before exiting. From the force of impact, inertia and the size of the media
web, particles are trapped within the filter. The result is that a very high efficiency is achieved, partic-
ularly with small diameter particles.

Extended surface filters have much higher resistance than electrostatic filters but they have a far

350/Appencfix II: Laminar Flow Systems

greater capacity for holding dust. As the filter traps dust, it increases in weight and airflow declines.
Generally a HEPA filter is not reused but discarded when, as a rule of thumb, the resistance or
“pressure drop” doubles. Extended surface filters are used in hospital surgery rooms as well as cul-
ture laboratories and nuclear facililties. Since its efficiency is somewhat dependent on the impact
velocity of the particle striking the media web, an appropriate fan must be matched with this type of
filter.

The Fan

When constructing a laminar flow hood, the filter size must be precisely fitted with a high pres-
sure fan. All fans are rated by the manufacturer according to the volume of air (CFM or cubic feet
per minute) they can push past materials of specified resistance. The type of high pressure fans
needed in a laminar flow hood are usually of the squirrel cage type (“furnace blowers”).

In turn, the resistance of all micron filters are measured in inches of static pressure at a certain
air speed. A standard resistance for a micron filter of this type is .75-1 .00 inches of static pressure.
Because extended surface filters have a high initial resistance, the housing must tightly hold the
HEPA filter so that impure air is not sucked into the exiting airstream.

To calculate the correct fan/filter combination, take the net CFM of the fan at the filter’s rated
level of resistance and divide that number by the square footage of the filter face. Ideally, that num-
ber will be 1 00 feet per minute, the optimum range for air velocity in laminar flow systems. An ex-
ample will more clearly illustrate this basic principle.

IF a micron filter measures 2 feet long by 2 feet high by 6 inches deep and has a static pressure
rating of 1 .0 inches of resistance, the fan required would have to be capable of pushing 400 CFM
at 1 inch of static pressure.

IF X = the desired net CFM of a fan at 1” S.P. and
Y = 4 square feet (the square footage of the filter face)

THEN X = 1 00 feet per minute x Y
X = 100 feet per minute x 4 square feet
X = 400 cubic feet per minute

This means that a fan capable of pushing 400 cubic feet per minute at 1 inch of static pressure
is needed to yield the optimum air velocity of 100 feet per minute. (Note that different filters have
different static pressure ratings and suggested CFM’s). In selecting a fan, it is best to choose one that
can deliver more than a 1 00 feet per minute air velocity. Install a solid state speed control to regu-
late the fan as needed.

As the filters become laden with particulates, the resistance increases and the airflow declines. If
the airflow falls below 20% of the suggested optimum, the 99.99% efficiency rating can not be
guaranteed. Filters of the size in the example above can hold four or more pounds of dust and
spores before needing replacement! With a few hours of use every week (the time most home culti-
vators spend conducting sterile transfers), the micron filter should last many years, depending of
course, on the ambient spore load in the laboratory.

Appendix II: Laminar Flow Systems/351

The life of a HEPA filter can be extended with the placement of prefilters to screen out coarse
particulates. Prefilters can be made of fiberglass media, the type commonly used for furnace filters,
or they can be composed of a thin open-celled foam. Prefilters of the latter type increase resistance
significantly whereas furna^jtype filters increase resistance only slightly. In this regard, furnace fil-
ters are well suited becawieOiiey are cheap (less than five dollars), readily available, and come in nu-

merous sizes.

Laminar Flow Designs

There are several types of laminar flow systems, each designed for specific applications. The
airflow in a biological safety cabinet, built for use with pathogenic organisms, is such that the worker
is not endangered if spores from a virulent organism became airborne. The air is drawn from the
work area into the hood and then up through micron filters and exited to the outside. Laminar flow
hoods for work with radioactive and toxic materials are similarly designed. Because of their intri-

Figure 241 Sterile room with ceiling composed of micron filters.

352/Appendix II: Laminar Flow Systems

cacy they are considerably more expensive than the kind needed for mushroom and plant culture.

Laminar flow systems for tissue culturists operate on a reverse principle of the one designed for
use with toxic substances. Air is forced through a micron filter to the work area, creating a positive
pressure sterile wind in which to conduct mycelial transfers. These types of hoods are perfect for
pouring media, maintaining pure mycelia and inoculating spawn containers. Since they greatly re-
duce the waste caused by contamination, their cost is soon offset by the savings realized. A laminar
flow hood is a low maintenance, affordable and appropriate technology for the serious home culti-
vator.

An alternative to building a laminar flow hood is the construction of a laminar flow wall or ceil-
ing. A laminar flow ceiling is preferable because the draft is directed downwards to the floor where it
exists through evenly placed pressure activated dampers. When a wall or ceiling is composed of mi-
cron filters, the air is usually drawn from the outside where the prefilters can be changed without en-
tering the sterile laboratory. Any contaminant spores tracked in on the shoes of workers are kept
close to the floor and is immediately swept away by the flow of sterile air. The atmosphere in a this
type of sterile room is fully exchanged 10-20 times per hour.

Foremost, tissue culturists are interested in preventing contamination from occurring, not from
spreading. They are concerned with creating sterile media and maintaining the purity of cultures. A
laminar flow hood is of little value in helping a cultivator isolate a colony of mushroom mycelium
away from, for instance, a green mold on a petri dish. The turbulence generated from the hood
would free thousands of spores, some of which would adhere to the surface of the sterile media, ger-
minate and produce more spores. In these cases, a laminar flow hood is best used as an air cleaner
prior to isolating a culture away from a contaminant. Several minutes after it has been turned off and
the air currents have settled, transfers can be made away from neighboring contaminants with little
danger of airborne spores.

Although sterile work can be conducted without a laminar flow system, they have become a
standard piece of equipment in professional spawn laboratories and increasingly in the sterile rooms
of many home cultivators.

Appendix III: Effects of Bacteria on Fruiting/353

A lthough mushrooms have been cultivated for more than two hundred years, little is known
about the biological processes of fruiting. For mushroom pinheads to form suddenly and then
to enlarge into towering mushrooms within only a few days represents a many hundred-fold multi-
plication in biomass. This ability to generate tissue so rapidly has few parallels in nature and has
been the subject of numerous scientific papers.

Mushrooms are in constant competition with organisms sharing the same habitat. Dung inhabit-
ing mushrooms in particular (like Psilocybe cubensis and Agaricus brunnescens) live in an envi-
ronment that teems with other microorganisms feeding on organic wastes and dead cell matter.
Dung is by nature a temporary substrate, decomposing completely in only a few weeks. Within this
short period of time there is a succession of dominant microorganisms, most notably fungi and bac-
teria. For a new mushroom colony to grow, its spores must fall, germinate, mate, form a substantial

Figure 242 Psilocybe cyanescens mycelium contaminated with
bacteria.

354/Appendix 111: Effects of Bacteria on Fruiting

I—— ’V ’ — —

mycelial network and then produce a specialized fruitbody. This series of events is made less likely
by poor weather conditions and/ or competing microorganisms. The brevity of the generative phase
in the mushroom life cycle suggests a highly advanced metabolic system, one that has evolved
despite its fiercely competitive environment.

The fact that Agaricus brunnescens fails to fruit on sterilized substrates has been well docu-
mented. It has been shown that if the casing layer is sterilized and applied to grain or compost,
mushrooms do not form. On the other hand, if the casing layer is only pasteurized or left untreated,
fruiting is unhindered. Obviously something in the peat based casing is essential to the fructification
process.

Past investigations have shown the significance of bacteria in mushroom growth. It should not
be surprising then to learn that some of these microorganisms are not harmful to the mushroom
plant, but beneficial. Under conditions of high humidity, C0 2 and acetone, bacterial populations
spiral. In a way not presently understood, some of these bacteria act as a trigger to fruiting. The
prevalence of bacteria on hyphae may explain why most dung dwelling mushrooms can be fruited
with comparative ease on basic enriched agar media while wood and soil inhabitors can not. The as-
sociation of these two organisms, a fungus and a bacterium, reflects a tacit agreement for mutual co-
existence, one perhaps negotiated by evolutionary necessity.

In 1956 Dr. Takashi Urayama first noted the stimulative influence of bacteria on the fruiting of
Psilocybe coprophila. (Actually he misidentified the mushroom species as P. panaeoliformis). In
that paper and ones soon thereafter (Urayama 1960, 1961 and 1967), he reported the isolation of
a bacterium he thought responsible for fruiting in not only Psilocybe “panaeoliformis” but also in
Agaricus brunnescens. He named that bacterium Bacillus psilocybe nom. prov. Apparently un-
aware of Urayama’s work, a German mycologist named Eger similarly isolated a bacterium stimula-
tive to pinhead formation. She first published her notes in 1959. For years this bacterium was
known as “Eger’s Bacterium” until Hayes (1969) identified the organism in question as
Pseudomonas putida. This identification set in motion other research projects whose conclusions
revealed a subtle but dynamic interplay between microflora in the casing layer and the mushroom
mycelium.

Mushroom mycelium releases several metabolites as it grows through a substrate, most impor
tantly C0 2 . Other compounds identified by researchers as metabolic waste products include ace-
tone, ethanol and ethylene. Upon casing, the release of volatile metabolites from the spawned com-
post or grain is drastically inhibited. The casing layer interferes with the free diffusion of acetone, and
hence its concentrations in the casing biosphere increase. Since Pseudomonas putida grows on
media whose sole carbon source is acetone or ethanol (2.5%), cultivators can adopt measures that
will enhance the levels of these Pseudomonas propagating compounds in the casing layer. Eger
first suggested a practical application for commercial cultivators:

“In order to prove our hypothesis, freshly prepared, moist casing casing soil of a commercial
mushroom plant should be incubated with acetone for several days apart from mushroom cultures.
If acetone has a stimulative effect on the microflora that induces fructification, soil treated with ace-
tone should allow earlier pinhead formation than control samples.” (Eger, 1972, pp. 723.)

Two years later Hayes and Nair (1974) noted that more bacteria flourish in wet casings placed

Appendix 111: Effects of Bacteria on Fruiting/355

on compost than in wet casing alone. Peak activity occurred ten days after application. Dry casings,
as one would expect, had significantly fewer bacteria. Continuing with this work, Hayes and Nair
showed that the addition of 5% spawned compost into the casing layer resulted in the largest in-
crease in P. putida populations, the most pinheads and the greatest overall yields.

Stanek (1974), a Czech mycologist, studied the bacteria associated directly with mushroom
mycelium, in the zone he called the ’’hyphosphere”. These hyphosphere bacteria differed from
other bacteria in that they were predominantly Gram-negative (as is Pseudomonas putida) and they
utilized nitrogenous compounds secreted by the mycelium. Both the growth of mycelia and bacteria
were stimulated by extracts of one another, suggesting a mutually enhancing relationship much like
the one between nitrogen fixing bacteria and the roots of many plants. Stanek further determined
that mycelium infected with bacteria grew more quickly through compost and would, therefore, give
mushroom mycelium a decided advantage over other competing microorganisms. From this
author’s experience (Stamets’) in the course of studying the hyphosphere of several Psilocybe
species, bacteria are not uncommon and may play a similarly beneficial role.

Not all strains of Pseudomonas putida cause pinheads to form in Agaricus brunnescens, nor
do all strains of mushrooms respond similarly to the presence of selected bacteria. The two proven
stimulative strains, ATCC #12633 and #17419, are deposited with the American Type Culture
Collection. Some strains of Pseudomonas putida have no effect whatsoever, while others are most
stimulative if the bacterial colonies are grown on a 2.5% acetone based liquid media (see Eger,
1972). After incubating for 10 days at 25 °C. in 30-40 ml. of nutrient broth, a density of
1 ,000,000 to 2,000,000 cells/milliliter is achieved. Ten milliliters of this concentrated solution is
recommended for each square meter of casing surface. (For ease of application, one milliliter of
concentrate can be diluted in 100 milliliters of sterilized water).

Eger, Hayes and Nair have demonstrated the stimulative effect of Pseudomonas putida. But
why Pseudomonas putida stimulates primordia formation is a question yet unanswered. Some be-
lieve its effect is indirect, removing chelating compounds that inhibit mushroom initiation. Others
(Fritsche, 1981; Visscher, 1981) suspect its influence is more direct and biologically oriented.

Pseudomonas putida is not the only microorganism implicated in the phenomenon of fruiting.
Park and Agnihorti (1 969) published a short note where they compared bacteria introduced to soils
that had been autoclaved, gamma sterilized and untreated. Three other bacteria ( Bacillus
megaterium, Arthrobacter terregens and Rhizobium meliloti ) stimulated abundant fruitbody forma-
tion and development on sterilized soils. (Interestingly, these same nitrogen fixing bacteria are pres-
ently being marketed to farmers for increasing crop production). In yet another study, Curto and
Favelli (1972) examined a gamut of microorganisms (bacteria, yeasts and microalgae) and their ef-
fect on potentiating yields. Again, Bacillus megaterium significantly increased mushroom forma-
tion. Even more remarkably Scenedesmus quadricauda (a common pond dwelling blue-green
alga) enhanced production by nearly 60% over and above the control. This alga seemed to have a
particularly influencial effect on the number of primordia generated on the first flush. Although as
exciting as these findings may at first appear, it must be noted that other researchers have not yet
confirmed the findings of Curto and Favelli. For reasons not presently understood, activated char-

356/ Appendix 111: Effects of Bacteria on Fruiting

coal mimics the primordia stimulating properties of Pseudomonas putida and other beneficial mi-
croorganisms. (See Chapter VIII). Its addition to unsterilized casings seems wholly unnecessary
considering the ease with which Agaricus brunnescens and Psilocybe cubensis form pinheads.
But, in sterilized casings or in casings applied to difficult to fruit species, the use of activated charcoal
and select bacteria gives the cultivator another means to promote fructification. Although many
studies have been published, work with fruiting potentiators is still in its in infancy. Specific mush-
room strains must be carefully matched with specific strains of potentiators. And the potentiators
themselves, while of value at fruiting, can be formidable competitors to sterile culture in the labora-
tory. Nevertheless, utilizing these benevolent microorganisms holds great promise for the future of
mushroom culture.

NOTE: Bacteria, if cultured, must be kept separate from the mushroom culture laboratory.
Pseudomonas and Bacillus grow well on standard 2% malt agar media.

Appendix IV: Extracts to Induce Primordia Formation/357

THE USE OF

A FORMATION

T he search for the biochemical means by which mushrooms fruit has been ongoing for years.

Several researchers have demonstrated the influence of hormones in regulating mushroom for-
mation and development. From this work, it is clear that no one mechanism, but many, cause the
phenomenon of fruiting.

Urayama (1972) found that live extracts from young buttons of Agaricus brunnescens and
from other species would induce pinhead initiation in a Marasmius species that otherwise failed to
fruit on a specified agar medium. He determined that this particular Marasmius failed to form fruit-
bodies on agar media that had a carbonmitrogen ratio of 1:10 with sucrose levels maintained at
1 %. Given the inability of pinheads to form at this Sucrose/Peptone ratio, he could introduce stan-
dardized cell free extracts of other mushroom species to gauge their effects. Species from which
crude extracts were taken were: Agaricus brunnescens, Lentinus edodes, Flammulina velutipes
and Pleurotus ostreatus. The extracts were performed by washing 200 grams of homogenized live
mushroom tissue (primordia less than 1 cm. tall) with four successive baths of 80% methanol. The
residue was discarded each time and the methanol solution allowed to evaporate, under a slight vac-
uum, until a dried filtrate remained. One gram of this crude extract was then immersed into 1 0 milli-
liters of water and applied in ’/ 0 th milliliter increments to each culture tube, except for the controls.

The results of Urayama’s work showed that each of the four fractionations induced primordia
formation provided aqueous methanol (80%) and only young mushrooms were used. Extracts from
older fru itbodies, especially that of Agaricus brunnescens and Lentinus edodes, had no effect what-
soever. Urayama tried other solvents to isolate the mysterious “fruiting hormone” and discovered
that it was soluble in water and not soluble in absolute methanol, chloroform or petroleum benzine.
He worked on his “Substance X”, as he liked to call it, for many years until his death in 1980.

Shiio et alia (1974) realized that young mushroom buttons contained high concentrations of
the fruiting hormones and applied this knowledge to the commercial cultivation of Flammulina
velutipes. Pieces of Flammulina velutipes primordia were immersed into sterile water and sprayed
over sawdust/bran beds. Not only were yields substantially increased by this crude procedure, but
initiation occurred much earlier, and the overall fruiting cycle was narrowed considerably. Clearly
these mycologists were on the road to discovering an important link in the biochemistry of fruiting.

Around the same time as the work of Shiio et alia, two other Japanese mycologists published
related studies (Uno & Ishikawa, 1971, 1 973) whereby pinheads of Coprinus formed if a “cell free
extract” from young mushrooms was added to the culture. They and others isolated the causal

358/Appendix IV: Extracts to Induce Primordia Formation

compounds— cyclic adenosine monophosphate (c AMP) and related enzymes. They further found
that light stimulated the production of c AMP in the mycelium of phototropic mushroom species.
Conversely, the absence of light in phototropic mushroom species resulted in no production of c
AMP.

Wood (1 979) tried to substantiate the findings of Uno and Ishikawa with Agaricus brunnescens
and failed. He could not induce primordia to form using c AMP. However, this fact does not bear
any significance on the importance of cyclic adenosine monophosphate in phototropic species
since A. brunnescens is a mushroom needing no light whatsoever for primordia formation and de-
velopment.

The question of how mushrooms fruit is not simple; nor will there be one answer explaining the
mechanisms in all species. What is apparent at this early stage of research is that photosensitive and
non-photosensitive species have developed different means for mushroom development. The infor-
mation most useful for home and commercial cultivators will come in the areas of yield enhance-
ment and the growing of exotic mushrooms on readily available, cheap materials. By good fortune,
this is one area of research that is not beyond the means of the innovative home cultivator.

Appendix V: Data Collection Records/359

S uccess in mushroom growing requires a consistent and repeatable methodology. Because
there are so many variables that affect the crop, careful record keeping is essential for good
management. With a data collection system, the cultivator can learn from mistakes and gain a
deeper understanding of the factors that influence healthy mushroom growth.

The following data collection records reflect years of mushroom growing experience and are
therefore quite detailed. Each cultivator must evaluate his or her particular circumstance to decide
which categories are most appropriate. In turn, these data sheets can be modified to meet an individ-
ual’s requirements.

360/Appendix V: Data Collection Records

Mushroom spprips'

SPAWN MAKING

Strain:

tissi ip-

spores:

Spawn media:
water

additives:

Sterilization time and temperature:
Inoculation date:

Date of Full Colonization:

Shaking schedule:

Observations: _

Temperature Chart

air temp.

Appendix V: Data Collection Records/361

COMPOST MAKING: Phase S

Compost formula

Ingredient

wet weight

\|fh 2 o

dry weight

% N

lbs. N

j Dj°

Percent Nitrogen:

Type of Straw: _____ Source of Horse manure:

Description:

structure:

color: - — —

age:

Date pre-composting started: —

Method of watering: _

Supplements added:

Number of turns: —

Total pre-composting time: _

Comments: __

362/Appendix V: Data Collection Records

Appendix V: Data Collection R ecords/363

COMPOST MAKING: Phase 51

°p compost temp. air temp.

364/Appendix V: Data Collection Records

Spawning Date:

Spawn type and amount:
Substrate description: _

pH:

Supplements:

SPAWN RUNNING

, Substrate density (lbs. dry wt/ft. 2 )

% H 2 0:

structure: .

Temperature Chart

. substrate temp. * air temp

ssssissasissssssisssssaHS

88888888838888888888888888
888888888S88888S888888888S
18888888888 888888 8 8883 88888

aBBflBiBBBflBflBBBBBBBBBflBflBBB

aBBBB&BBBBBBBBBBBflBflBflflBflBK

■BBfliaaa ibbbbbbi bum ■■

BBflBflBBBBBBflBflBBBBBaBBBflBB

Fan speed

% Fresh air

Heat/thermo
stat setting

Humidity

laaiwnsiB Baa

BBBBainai

888888888^1

B— —BlitB

Appendix V: Data Collection Records/365

Casing formula:

CASING: CASE RUNNING

— pH:

% moisture at application:

Substrate supplementation:

Scratching: _ , — _

depth:

Patching:

substrate temp. — air temp.

HMBiBBHHHIl

■mBhBMI

Date/time

Fan speed

% Fresh air

FI eat/thermo
stat setting

C0 2

Humidity

Watering

Light

Appendix V: Data Collection Records/ 367

CROPPING

Evenness of pin set: ,

% of surface pinned: — —

Date of first harvest:

Temperature Chart

substrate temp. * — - air temp.

Days

Contaminants encountered:

Total time (filling to emptying):

Appendix VI: Analyses of Materials/369

DRY ROUGHAGES OF FIBROUS MATERIALS

Total dry Protein

Fat

Fiber

N-free

Total

Calcium

Phos-

Nitro-

Potas-

matter

extract

minerals

phorus

gen

sium

Per a.

Per ct.

Alfalfa hay, all analyses

90.5

14.8

2.0

28.9

36.6

8.2

1.47

0.24

2.37

2.05

Alfalfa hay, very leafy (less than 25% fiber)

90.5

17.2

2.6

22.6

39.4

8.7

1.73

0.25

2.75

2.01

Alfalfa hay, leafy (25-28% fiber)

90.5

15.8

2.2

27.4

36.6

8.5

1.50

0.24

2.53

2.01

Alfalfa hay, stemmy (over 34% fiber)

90.5

12.1

1.4

36.0

33.4

7.6

1.10

0.18

1.94

1.68

Alfalfa hay, before bloom

90.5

19.0

2.7

22.6

36.7

9.5

2.22

0.33

3.04

2.14

Alfalfa hay, past bloom

90.5

12.8

2.1

31.9

36.2

7.5

—

2.05

Alfalfa hay, brown

87.9

17.3

1.6

24.5

35.1

9.4

1.37

0.26

2.77

—

Alfalfa hay, black

83.1

17.5

1.5

29.1

25.3

9.7

—

2.80

—

Alfalfa leaf meal

92.3

21.2

2.8

16.6

39.7

12.0

1.69

0.25

3.39

—

Alfalfa leaves

90.5

22.3

3.0

14.2

40.5

10.5

2.22

0.24

3.57

2.06

Alfalfa meal

92.7

16.1

2.2

27.1

38.2

9.1

1.32

0.19

2.58

1.91

Alfalfa stem meal

91.0

11.5

1.3

36.3

34.8

7.1

—

1.84

—

Alfalfa straw

92.6

8.8

1.5

40.4

35.1

6.8

—

0.13

1.41

—

Alfalfa and bromegrass hay

89.3

12.4

2.0

28.6

38.1

8.2

0.74

0.24

1.98

2.18

Alfalfa and timothy hay

89.8

11.1

2.2

29.5

40.3

6.7

0.81

0.21

1.78

Alfilaria, dry (Erodium cicutarium)

89.2

10.9

2.9

23.4

40.2

11.8

1.57

0.41

1.74

Alfilaria, dry, mature

89.0

3.5

1.5

31.4

44.1

8.5

—

0.56

—

Atlas sorghum stover

85.0

4.0

2.0

27.9

44.2

6.9

0.34

0.09

0.64

—

Barley hay

90.8

7.3

2.0

25.4

49.3

6.8

0.26

0.23

1.17

1.35

Barley straw

90.0

3.7

1.6

37.7

41 .0

6.0

0.32

0.1 1

0.59

1.33

Bean hay, mung

90.3

9.8

2.2

24.0

46.6

7.7

—

1.57

—

Bean hay, tepary

90.0

17.1

2.9

24.8

34.7

10.5

—

2.74

—

Bean pods, field, dry

91.8

7.1

1.0

34.8

45.0

3.9

0.78

0.10

1.14

2.02

Bean straw, field

89.1

6.1

1.4

40.1

34.1

7.4

1.67

0.13

0.98

1.02

Beggarweed hay

90.9

15.2

2.3

28.4

37.2

7.8

1.05

0.27

2.43

2.32

Bent grass hay, Colonial

88.5

6.6

3.0

29.5

42.8

6.6

—

0.18

1.06

1.42

Bermuda grass hay

90.6

7.2

1.8

25.9

48.7

7.0

0.37

0.19

1.15

1.42

Bermuda grass hay, poor

90.0

5.8

0.9

38.8

37.7

6.8

—

0.93

—

Berseem hay, or Egyptian clover

91.7

13.4

2.7

21.0

42.7

11.9

3.27

0.28

2.14

2.05

Birdsfoot trefoil hay

90.5

13.8

2.1

27.5

41.2

5.9

1.13

0.22

2.35

1.52

Black grass hay (Juncus Cerardi)

89.7

7.5

2.5

25.1

47.3

7.3

—

0.09

1.20

1.56

Bluegrass hay, Canada

89.3

6.6

2.3

28.2

46.4

5.8

0.20

1 .Ub

1 .94

Material

Bluegrass hay, Kentucky, all analyses
Bluegrass hay, Kentucky, in seed
Bluegrass hay, native western

Bluejoint hay (Calamagrostis Canadensis)
Bluestem hay (Andropogon, spp.)
Bromegrass hay, all analyses
Bromegrass hay, before bloom
Broom c orn stover

Buckwheat hulls
Buckwheat straw

Buffalo grass hay (Bulbilis dactyloides)
Bunchgrass hay, misc. varieties

Ca rpet grass hay

Cat-tail, or tule hay (Typha angustifolia)

Cereals, young, dehydrated

Chess, or cheat hay (Bromus, spp.)

Clover hay, alsike, all analyses

Clover hay, alsike, in bloom

Clover hay, Alyce
Clover hay, bur
Clover hay, crimson
Clover hay, Ladino

Clover, Lading, and grass hay

Clover hay, mammoth red

Clover hay, red, all analyses

Clover hay, red, leafy (less than 25% fiber)

Clover hay, red, stemmy (over 31% fiber)

Clov er hay, red, before bloom

Clover hay, red, early to full bloom

Clover hay, red, second cutting

Clover hay, sweet, first year

Clover hay, sweet, second year

Clover hay, white

Clover leaves, sweet

Clover stems, sweet

Clover straw, crimson

Clover and mixed grassy, high in clover

Clover and timothy hay, 30 to 50% clover

Corn cobs, ground
Corn fodder, well-eared, very dry
(from barn or in arid districts)

Corn fodder, high in water
Corn fodder, sweet corn
Corn husks, dried

Corn leaves, dried
Corn stalks, dried

Total dry Protein
matter

Per ct. Per cl.

89.4 8.2

87.3 5.5

91.9 11.2

Per ct. Per ct.

N-free Total Calcium Phos- Nitro- Potas-

extract minerals phorus gen sium

Per ct. Per ct. Per ct. Per ct. Per ct. Per ct.

92.8 24.5

89.0 10.9

92.1 18.4
89.5 14.2
88.0 19.4

2.8 29.8 42.1
2.5 31.0 41.9

3.0 29.8 39.9

2.3 32.9 39.6

2.2 30.2 43.4

2.1 28.4 39.5

2.3 24.6 37.9

1.8 36.8 42.4

1.0 42.9 40.1
1.0 36.2 38.8

6.5 0.46 0.32 1.31 1.73

6.4 0.23 0.20 0.88 1.48

8.0 — — 1 .79 —

6.5 - -

5.4 - -

8.2 0.20 0.28

1.15 -

0.86 -
1.58 2.35

1.6 0.26 0.02 0.48 0.27
8.3 1.24 0.04 0.69 2.00

1.8 23.8 46.2 10.1 0.70 0.13 1.09

2.0 30.4 44.1 9.4 - - 0.93

2.2 31.8 40.9 10.2 - - 1.12

92.2 26.6

90.4 2.3

1.7 30.8 44.3

4.7 16.1 33.1
2.1 29.2 46 1

2.1 27.0 39.9

3.2 26.9 37.7

1.6 35.4 35.5
2.9 22.9 37.8

2.2 27.4 37.0

3.2 20.7 34.9

2.2 20.7 41.7

3.4 29.2 37.0

2.6 27.2 40.1

3.1 23.6 40.8

2.1 34.1 36.0

3.6 18.0 41.1

3.5 26.1 39.7

2.9 24.5 40.4

2.5 24.6 39.7

1.9 30.2 37.6

2.4 22.5 40.9

3.2 9.5 41.9

1.1 38.0 35.6

1.5 38.8 32.9

2.7 28.8 42.2

2.2 30.3 41.2

0.4 32.1 54.0

2.2 27.1 47.6

1.4 16.7 34.2

1.8 26.4 41.3
0.9 28.2 49.6

1.9 23.9 42.6

1.5 28.0 43.3

8.2 -

14.4 0.66 0.46
7.4 0.29 0.25
7.8 1.15 0.23
7.8 1.32 0.25

1.32 0.45
1.23 0.24
1.32 0.29
1.05 0.26

0.93 -

3.92 -

1.10 1.47

1.94 2.44
2.14 2.27

1.74 -

2.94 2.96
2.27 2.79

3.10 2.78

6.7 - 0.24 1.87 -

6 4 1.35 0.19 1.89 1.43

7.2 - - 2.14 -

5.9 0.99 0.15 1.62 1.77

7.1 1.69 0.28 2.93 2.26

6.3 1.47 0.22 2.00 1.73

6.9 - - 2.14 -

8.5 1.37 0.26 2.64 1.57

7.5 1.25 0.23 2.16 1.78

7.8 1.16 0.24 2.30 1.66

11.0 - - 4.26 -

7.4 - — 1.70 -

7.0 - — 120 —

6.2 0.90 0.19 1.54 1.46

5.8 0.68 0.20 1.38 1.47

]~6 - 0.02 0.37~Ch37~

6.4 0.24 0.16
3.6 0.16 0.11

1.25 0.82
0.77 0.55

9 0 - 0.17 1.47 0.98

2.9 0.15 0.12 0.54 0.55

6.7 0.29 0.10 1.23 0.36

5.3 0.25 0.09 0.75 0.50

Appendix VI: Analyses of Materials/371

Material

Total dry Protein
matter

Pei cl. Per ct.

Fat

Per ct.

Fiber
Per ct.

N-free
extract
Per ct.

Total
minerals
Per ct.

Calcium
Per ct.

Phos-
phorus
Per ct.

Nitro-
gen
Per ct.

Potas-
sium
Per ct.

Corn stover (ears removed), very dry

90.6

5.9

1.6

30.8

4.65

5.8

0.29

0.05

0.94

0.67

Corn stover, high in water

59.0

3.9

1.0

20.1

30.2

3.8

0.19

0.04

0.62

0.44

Corn tops, dried

82.1

5.6

1.5

27.4

42.0

5.6

—

0.90

—

Cotton bolls, dried

90.8

8.7

2.4

30.8

42.0

6.9

0.61

0.09

1.39

3.18

Cotton leaves, dried

91.7

15.3

6.8

10.3

43.5

15.8

4.58

0.18

2.45

1.36

Cotton stems, dried

92.4

5.8

0.9

44.0

37.5

4.2

—

0.93

Cottonseed hulls

90.7

3.9

0.9

46.1

37.2

2.6

0.14

0.07

0.62

0.87

Cottonseed hull bran

91.0

3.4

0.9

37.2

46.7

2.8

—

0.54

—

Cowpea hay, all analyses

90.4

18.6

2.6

23.3

34.6

11.3

1.37

0.29

2.98

1.51

Cowpea hay, in bloom to early pod

89.9

18.1

3.2

21.8

36.7

10.1

—

2.90

Cowpea hay, ripe

90.0

10.1

2.5

29.2

41.8

6.4

—

1.62

—

Cowpea straw

91.5

6.8

1.2

44.5

33.6

5.4

—

1.09

Crabgrass hay

90.5

8.0

2.4

28.7

42.9

8.5

—

1.28

—

Durra fodder

89.9

6.4

2.8

24.1

51.4

5.2

1.02

Emmer hay

90.0

.97

2.0

32.8

36.4

9.1

, —

—

1.55

—

Fescue hay, meadow

89.2

7.0

1.9

30.3

43.2

6.8

—

0.20

1.12

1.43

Fescue hay, native western (Festuca, spp.)

90.0

8.5

2.0

31.0

42.8

5.7

—

1.36

—

Feterita fodder, very dry

88.0

8.0

2.1

18.7

51.5

7.7

0.30

0.21

1.28

—

Feterita stover

86.3

5.2

1.7

29.2

41.9

8.3

0.83

Flat pea hay

92.3

22.7

3.2

27.7

32.0

6.7

—

0.30

3.63

2.02

Flax plant by product

91.9

6.4

2.1

44.4

33.1

5.9

—

1.02

—

Flax straw

92.8

7.2

3.2

42.5

32.9

7.0

0.48

0.07

1.15

0.73

Fowl meadow grass hay

87.4

8.7

2.3

29.7

39.5

7.2

—

1.39

—

Furze, dried

94.5

11.6

2.0

38.5

35.5

7.0

1.86

Gama grass hay (Tripsacum dactyloides)

88.2

6.7

1.8

30.4

43.1

6.2

—

1.07

—

Grama grass hay (Bouteloua, spp.)

89.8

5.8

1.6

28.9

45.6

7.9

0.34

0.18

0.93

—

Grass hay, mixed, eastern states, good quality

89.0

7.0

2.5

30.9

43.1

5.5

0.48

0.21

1.12

1.20

Grass hay, mixed, second cutting

89.0

12.3

3.3

24.8

41.7

6.9

0.79

0.31

1.97

1.15

Grass straw

85.0

4.5

2.0

35.0

37.8

5.7

0.72

Guar hay (Cyamposis psoraloides)

90.7

16.5

1.3

19.3

41.2

12.4

—

2.64

—

Hegari fodder

86.0

6.2

1.7

18.1

52.5

7.5

0.27

0.16

0.99

Ffegari stover

87.0

5.6

1.8

28.0

41.7

9.9

0.33

0.08

0.90

—

Flops, spent, dried

93.8

23.0

3.6

24.5

37.4

5.3

—

3.68

—

Florse bean hay

91.5

13.4

0.8

22.0

49.8

5.5

—

2.14

Florse bean straw

87.9

8.6

1.4

36.4

33.1

8.4

— .

—

1.38

— .

Hyacinth bean hay (Dilichos lablab)

90.2

14.8

1.4

33.6

6.8

—

2.37

—

Johnson grass hay

90.1

6.5

2.1

30.4

43.7

7.4

0.87

0.26

1.04

1.22

June grass hay, western (Koeleria cristata)

88.3

8.1

2.5

30.4

40.5

6.8

—

1.30

—

Kafir fodder, very dry

90.0

8.7

2.6

25.5

44.2

9.0

0.35

0.18

1.39

1.53

Kafir fodder, high in water

71.7

6.5

2.7

21.6

37.6

3.3

0.28

0.14

1.04

1.23

Kafir stover, very dry

90.0

5.5

1.8

29.5

44.3

8.9

0.54

0.09

0.88

—

Kafir stover, high in water

72.7

3.8

1.3

23.7

36.6

7.3

0.44

0.07

0.61

— ,

Koahaole forage, dried

88.7

12.7

1.9

29.8

39.2

5.1

—

2.03

—

Kochia scoparia hay

90.0

1 1.4

1.5

23.6

40.7

12.8

1.82

—

Kudzu hay

89.0

15.9

2.5

28.6

35.1

6.9

2.78

0.21

2.54

—

Lespedeza hay, annual, all analyses

89.2

12.7

2.4

26.7

42.2

5.2

0.98

0.18

2.03

0.91

372/Appendix VI: Analyses of Materials

Total dry Protein Fat Fiber
matter

Per ct. Per ct. Per ct. Per ct.

N-free Total Calcium Phos- Nitro- Pon

extract minerals phorus gen siu

Per ct. Per ct. Per ct. Per ct. Per ct. Per

Lespedeza hay, annual, before bloom
Lespedeza h ay, annual, in bloom

Lespedeza hay, annual, after bloom
Lespedeza hay, perennial
Lespedeza leaves, annual
Lespedeza stems, annual
Lesp edeza straw

Lovegrass hay, weeping

Marsh or swamp hay, good quality

Millet hay, foxtail varieties

Millet hay, hog millet, or proso

Millet hay, Japanese

Millet hay, pearl, or cat-tail
Millet straw
Milo fodder
Milo stover

Mi nt hay

Mixed hay, good, less than 30% legumes
Mixed hay, good, more than 30% legumes
Mixed hay, cut very early
Napier grass hay

Natal grass hay

Native hay, western mt. states, good quality
Native hay, western mt. states, mature and
weathered

Needle grass hay (Stipa, spp.)

Oak leaves, live oak, dried

Oat chaff

Oat hay

Oat hay, wild (Avena fatua)

Oat hulls
Oat straw

Oa t grass hay, tall

Orchard grass hay, early-cut
Picnic grass hay (Panicum, spp.)

Para grass hay

Pasture grasses and clovers, mixed, from
closely grazed, fertile pasture, dried
(northern states)

Pasture grasses, mixed, from poor to fair
pasture, befo re heading out, dried

Pasture grass, western plains, growing, dried
Pasture grass, western plains, mature, dried
Pasture grass, western plains, mature and
weathered

86.8 8.3

2.7 22.7 43.0

1.8 26.5 42.7

1.9 32.6 38.6

1.7 26.5 42.7

2.9 19.7 43.1

1.0 38.5 37.7
2.3 29.2 47.1

2.8 30.9 43.4
2.3 28.2 44.3
2.7 25.3 44.7

2.2 23.9 47.6

1.6 27.7 40.8

1.7 33.0 36.8
1.6 37.5 41.6

3.3 21.9 48.4

1.1 29.1 48.1

2.1 20.3 45.6

6.4 1.04 0.19 2.29 1.06

5.1 1.02 0.18 2.08 0.94

4.5 0.90 0.15 1.84 0.82

4.9 0.92 0.22 2.11 0.98

6.4 1.30 0.20 2.74 0.92

3.7 0.64 0.13 1.33 0.89

6.7 0.29 0.16 1.31

7.3 - - 1.49

8.4 0.20 - 1.33

1.49 -

1.33 2.10

6.9 0.35 0.18 1.28 -

9.5 0.58 0.11 0.51 -

7.6 1.51 0.19 2.03 -

1.8 30.7 41.8 5.4 0.61

1.33 1.47

1.9 28.1 42.8

2.7 25.3 39.4

1.8 34.0 34.6
1.8 36.8 39.2

2.1 29.8 43.2

6.0 0.90 0.19 1.47

9.3 - - 2.13

10.5 - - 1.31

5.0 0.45 0.29 1.18

6.8 0.39 0.12 1.30

90.0 3.9

88.1 7.2

93.8 9.3

91.8 5.9

88.6 7.7

92.1 8.3

90.2 4.6

1.4 33.6 43.6 7.5 -

2.0 30.8 41.9 6.2 -

2.7 29.9 45.3 6.6 -

2.4 25.7 46.3 11.5 0.80

2.7 28.1 42.2 6.9 0.21

2.6 32.5 44.0 6.8 0.22

1.3 29.7 50.8 6.5 0.20-

2.2 36.1 41.0 6.3 O.fl^l

2.4 30.1 42.7 6.0

2.9 30.5 40.7 6.8 0.1?

2.3 29.5 44.9 7.1 -

0.9 33.6 44.5 6.6 0.35

- 0.62 -

- 1.15 -

- 1.49 -

0.30 0.94 0.86

0.19 1.31 0.83

, £i| 0.78 0.48
K|1jf[)o0.66 1 -35
120 1.36

1.23 1.61

- 1.33 -

0.35 0.74 1.44

90.0 20.3 3.6 19.7 38.7 7.7 0.58 0.32 3.25 2.18

90.0 14.1 2.3 19.4 43.2 11.0 0.41 0.12 2.26 0.74

90.0 11.6 2.5 28.0 40.2 7.7 0.37 0.24 1.86 -

90.0 4.6 2.3 31.9 45.3 5.9 0.34 0.14 0.74 —

90.0. 3.3 1.8 34.1 44.5 6.3 0.33 0.09 0.53 -

Appendix VI: Analyses of Materials/373

Total dry

Protein

Fat

Fiber

N-free

Total

Calcium

Phos-

Nitro-

Potas-

Material

matter

extract

minerals

phorus

gen

sium

Per ct.

Pasture grass and other forage on western

mt. ranges, spring, dried

90.0

17.0

3.1

14.0

49.1

6.8

1.21

0.38

2.72

—

Pasture grass and other forage on western

mt. ranges, autumn, dried

90.0

8.8

4.3

17.4

51.4

8.1

—

1.41

—

Pea hay, field

89.3

14.9

3.3

24.3

39.1

7.7

1.22

0.25

2.38

1.25

Pea straw, field

90.2

6.1

1.6

33.1

44.0

5.4

—

0.10

0.98

1.08

Pea-and-oat hay

89.1

12.1

2.9

27.2

39.1

7.8

0.72

0.22

1.94

1.04

Peanut hay, without nuts

90.7

10.1

3.3

23.4

44.2

9.7

1.12

0.13

1.62

1.25

Peanut hay, with nuts

92.0

13.4

12.6

23.0

34.9

8.1

1.13

0.15

2.14

0.85

Peanut hay, mowed

91.4

10.6

5.1

23.8

42.2

9.7

—

1.70

—

Peanut hulls, with a few nuts

92.3

6 7

1.2

60.3

19.7

4.4

0.30

0.07

1.07

0.82

Peavine hay, from pea-cannery vines,

sun-cured

86.3

11.9

2.4

23.0

42.2

6.8

1.48

0.16

1.90

—

Prairie hay, western, good quality

90.7

.57

2.3

30.4

44.9

7.4

0.36

0.18

0.91

—

Prairie hay, western, mature

91.7

3.8

2.4

31.9

47.1

6.5

0.28

0.09

0.61

0.49

Quack grass hay

89.0

6.9

1.9

34.5

38.8

6.9

—

1.10

—

Ramie meal

92.2

19.2

3.8

20.1

35.9

13.2

4.32

0.22

3.07

—

Red top hay

91.0

7.2

2.3

29.3

45.3

6.9

0.33

0.23

1.15

1.93

Reed canary grass hay

91.1

7.7

2.3

29.2

44.3

7.6

0.33

0.16

1.23

—

Rescue grass hay

90.2

9.8

3.2

24.6

44.5

8.1

—

1.57

—

Rhodes grass hay

89.0

5.7

1.3

31.7

41.8

8.5

0.35

0.27

0.91

1.18

Rice hulls

92.0

3.0

0.8

40.7

28.4

19.1

0.08

0.48

0.31

Rice straw

92.5

3.9

1.4

33.5

39.2

14.5

0.19

0.07

0.62

1.22

Rush hay. western (Juncus, spp.)

90.0

9.4

1.8

29.2

44.2

5.4

—

1.50

—

Russian thistle hay

87.5

8.9

1.6

26.9

37.4

12.7

—

1.42

—

Rye grass hay, Italian

88.6

8.1

1.9

27.8

43.3

7.5

—

0.24

1.30

1.00

Rye grass hay, perennial

88.0

9.2

3.1

24.2

43.4

8.1

—

0.24

1.47

1.25

Rye grass hay, native western

87.4

7.8

2.1

33.5

37.6

6.4

—

1.25

—

Rye hay

91.3

6.7

2.1

36.5

41.0

5.0

—

0.18

1.07

1.05

Rye straw

92.8

3.5

1.2

38.7

45.9

3.5

0.26

0.09

0.56

0.90

Salt bushes, dried

93.5

13.8

1.6

22.1

38.8

17.2

1.88

0.1 1

2.21

4.69

Salt grass hay, misc. var.

90.0

8.1

1.8

28.8

39.5

11.8

—

1.30

—

Sanfoin hay (Onobrychis viciaefolia)

84.1

10.5

2.6

19.7

44.2

7.1

—

1.68

—

Seaweed, dried (Fucus, spp.)

88.7

5.2

4.2

9.4

53.6

16.3

—

0.83

Seaweed, dried [Laminaria, spp.)

83.7

11.4

1.1

8.6

45.8

16.8

—

1.82

—

Sedge hay, eastern (Carex, spp.)

90.7

6.1

1.7

29.2

46.3

7.4

—

0.98

—

Sedge hay, western (Carex, spp.)

90.6

10.1

2.4

27.3

44.0

6.8

0.60

0.24

1.62

—

Seradella hay

89.0

16.4

3.2

29.8

32.0

7.6

—

0.33

2.62

1.25

Sorghum bagasse, dried

89.3

3.1

1.4

31.3

50.0

3.5

—

0.50

—

Sorghum fodder, sweet, dry

88.8

6.2

2.4

25.0

48.1

7.1

0.34

0.12

0.99

1.29

Sorghum fodder, sweet, high in water

65.7

4.5

2.4

16.6

37.6

4.6

0.25

0.09

0.72

0.96

Soybean hay, good, all analyses

88.0

14.4

3.3

27.5

35.8

7.0

0.94

0.24

2.30

0.82

Soybean hay, in bloom or before

88.0

16.7

3.3

20.6

37.8

9.6

1.53

0.27

2.67

0.86

Soybean hay, seed developing

88.0

14.6

2.4

27.2

36.5

7.3

1.35

0.25

2.34

0.78

Soybean hay, seed nearly ripe

88.0

15.2

6.6

24.0

38.2

4.0

0.86

0.32

2.43

0.81

Soybean hay, poor qualify, weathered

89.0

9.2

1.2

41.0

30.4

7.2

0.94

—

1.47

— - - — —

Material

Total dry Protein
matter

Per ct. Per ct. F

Fat

5 er ct.

Fiber
Per ct.

N-free Total Calcium

extract minerals

Per ct. Per ct. Per ct.

Phos-
phorus
Per ct.

Nitro-
gen
Per ct.

Potas-
sium
Per ct.

Soybean straw

Soybean and Sudan grass hay, chiefly Sudan

Spanish moss, dried

Sudan qrass hay, all analyses

88.8

89.0

89.2

89.3

4.0
7.4

5.0
8.8

1.1

2.2

2.4

1.6

41.1

31.1
26.6
27.9

37.5

43.4

47.7

42.9

5.1
4.9
7.5

8.1

0.36

0.13

0.04

0.26

0.64

1.18

0.80

1.41

0.62

0.46

1.30

Sudan grass hay, before bloom

89.6

11.2

1.5

26.1

41.3

9.5

0.41

0.26

1.79

—

Sudan grass hay, in bloom
Sudan grass hay, in seed
Sudan grass, young, dehydrated

89.2

89.5

88.0

8.4

6.8

14.5

1.5

1.6
2.5

30.7

29.9

20.4

41.8

44.4

41.2

618

6.8

9.4

0.27

0.52

0.19

0.39

1 .34
1.09
2.32

Sudan qrass straw

90.4

7.1

1.5

33.0

42.3

675

—

1.14

—

Sugar cane fodder, Japanese, dried

89.0

1.3

1.8

19.7

64.3

1.9

0.32

0.14

0.21

0.58

Sugar cane bagasse, dried

95.5

1.1

0.4

49.6

42.0

2.4

—

0.18

—

Sugar cane pulp, dried

93.8

1.7

0.6

45.6

42.2

3.7

—

0.27

—

Sweet potato vine, dried

90.7

12.6

3.3

19.1

45.5

10.2

— *

—

2.02

—

Teosinte fodder, dried

89.4

9.1

1.9

26.5

41.7

10.3

—

0.17

1 .46

0.88

Timothy hay, all analyses

89.0

6.5

2.4

30.2

45.0

4.9

0.23

0.20

1.04

1.50

Timothy hay, before bloom

89.0

9.7

2.7

27.4

42.7

675

—

1 .55

—

Timothy, full bloom

89.0

6.4

2.5

30.4

44.8

4.9

0.23

0.20

1.02

1 .50

Timothy hay, in bloom, nitrogen fertilized

89.0

9.7

2.1

31.6

42.6

3.9

0.40

0.21

1.41

Timothy hay, late seed

89.0

5.3

2.3

31.0

45.9

4.5

0.14

0.15

0.85

1 .41

Timothy hay, in bloom, dehydrated

89.0

7.7

2.3

28.3

45.5

5.2

1.23

—

Timothy hay, second cutting

88.7

15.0

4.6

25.4

36T5

7.2

—

2.40

—

Timothy and clover hay, one-fourth clover

88.8

7.8

2.4

29.5

43.8

5.3

0.51

0.20

1 .25

1 .48

Velvet bean hay

92.8

16.4

3.1

27.5

38.4

7.4

—

0.24

2.62

2.20

Vetch hay, common

89.0

13.3

1.1

25.2

32.2

6.2

1.18

0.32

2.13

2.22

Vetch hay, hairy

88.0

19.3

2.6

24.5

33.1

8.5

1.13

0.32

3.09

1.96

Vetch-and-oat hay, over half vetch

87.6

11.9

2.7

27.3

37.5

8.2

0.76

0.27

1.90

1 .51

Vetch-and-wheat hay, cut early

90.0

15.4

2.2

28.8

36.4

7.2

—

2.46

—

Wheat chaff

90.0

4.4

1.5

29.4

47.1

7.6

0.21

0.14

0.70

0.50

Wheat hay

90.4

6.1

1.8

26.1

50.0

6.4

0.14

0.18

0.98

1.47

Wheat straw

92.5

3.9

1.5

36.9

41.9

8.3

0.21

0.07

0.62

0.79

Wheat grass hay, crested, cut early

90.0

9.2

2.0

32.2

40.2

6.4

—

1.47

—

Wheat grass hay, slender

90.0

8.0

2.1

32.2

41.0

6.7

0.30

0.24

1.28

2.41

Winter fat, or white sage, dried (Eurotia lanata)

92.6

12.9

1.9

27.4

40.8

9.6

—

2.06

Wire qrass hay, southern (Aristida, spp.)

90.0

5.5

1.4

31.8

47.9

3.4

0. 1 5

0.14

0.88

—

Wire grass hay, western (Aristida, spp.) 90.0 6.4 1.3 34.1

Yucca, or beargrass, dried 92.6 6.6 2.2 38.6

CONCENTRATES

41.0

38.3

7.2

6.9

—

1.02

1.06

—

Material

Total dry Protein
matter

Per ct. Per ct.

Fat

Per ct.

Fiber
Per ct.

N-free
extract
Per ct.

Total
mineral*
Per ct.

Calciurr
Per ct.

i Phos-
phorus
Per ct.

Nitro-
gen
Per ct.

Potas-
sium
Per ct.

Acorns, whole (red oak)

Acorns, whole (white and post oaks)
Alfalfa-molasses feed
Alfalfa seed

50.0

86.0
88.3

3.2

2.7

11.4

33.2

10.7

9.9

25.0

1.2 -

- 0.51 -

3.0

9.3

33.7

1.3 -

- 0.43 -

1.2

18.5

46.2

8.7 -

- 1 .82 -

10.6

8.1

32.0

4.4 —

- 5.31 -

Appendix VI: Analyses of Materials/375

Total dry Protein

Fat

Fiber

N-free

Total

Calcium

Phos-

Nifro-

Potas-

matter

extract

minerals

phorus

gen

sium

Per ct.

Alfalfa seed screenings

90.3

31.1

9.9

li.i

33.1

5.1

4.98

Apple-pectin pulp, dried

91.2

7.0

7.3

24.2

49.4

3.3

—

1.12

—

Apple-pectin pulp, wet

16.7

1.5

0.9

5.8

7.9

0.6

—

0.24

—

89.4

4.5

5.0

15.6

62.1

2.2

0.10

0.09

0.72

0.43

Apple pomace, wet

21.1

1.3

3.7

13.9

0.9

0.02

0.21

0.10

Atlas sorghum grain

89.1

1 1.3

3.3

2.0

/U.b

1.9

—

1.81

—

Atlas sorghum head chops 88.0

Avocado oil meal 91.4

Babassu oil meal 92.8

Bakery waste, dried (high in fat) 91.6

Barley, common, not including Pacific Coast
states 89.4

9.5

18.6

24.2

10.9

12.7

2.8

1.1

6.8

13.7

1.9

10.7

17.6

12.0

0.7

5.4

60.2

36.0

44.6

64.7

66.6

4.8
18.1

5.2

1.6

2.8

0.13

0.06

0.71

0.37

1.52

2.98

3.87

1.74

2.03

0.49

Barley, Pacific Coast states

89.8

8.7

1.9

5.7

70.9

2.6

—

1.39

Barley, light weight

89.1

12.1

2.1

7.4

64.3

3.2

1.94

—

Barley, hull-less, or bald

90.2

11.6

2.0

2.4

72.1

2.1

—

1 .86

—

Barley feed, high grade

90.3

13.5

3.5

8.7

60.5

4.1

0.03

0.40

2.16

0.60

Barley feed, low grade

92.0

12.3

3.45

14.7

56.2

b.3

—

1.97

“

Barley, malted

93.4

12.7

2.1

5.4

70.9

2.3

0.06

0.42

2.03

0.37

Barley screenings
Beans, field, or navy

88.6

90.0

1 1.6
22.9

2.7

1.4

9.1

4.2

61.3

57.3

3.9

4.2

0.15

0.57

1.86

3.66

1.27

89.0

23.0

1.2

4.1

56.8

3.9

—

3.68

—

Beans, lima

89.7

21.2

1.1

4.7

58.2

4.5

0.09

0.37

3.39

1.70

Beans, mung

90.2

23.3

1.0

3.5

58.5

3.9

—

3.73

Beans, pinto

89.9

22.5

1.2

4.1

57.7

4.4

—

3.60

—

Beans, tepary
Beechnuts

90.5

91.4

22.2

15.0

1.4

30.6

3.4

15.0

59.3

27.5

4.2

3.3

0.58

0.30

3.56

2.40

0.62

Beef scraps

94.5

55.6

10.9

1.2

0.5

26.3

—

8.90

—

Beet pulp, dried

90.1

9.2

0.5

19.8

57.2

3.4

0.67

0.08

1.47

0.18

Beet pulp, molasses, dried

91.9

10.7

0.7

16.0

59.4

5.1

0.62

0.09

1.71

1 .63

Beet pulp, wet

11.6

1.5

0.3

4.0

5.3

0.5

0.09

0.01

0.24

0.02

Beet pulp, wet, pressed

14.2

1.4

0.4

4.6

7.1

0.7

—

0.22

—

Blood flour, or soluble blood meal

92.2

84.7

1.0

1.1

0.7

4.7

0.68

U.5U

1 3.5b

—

Blood meal

91.8

84.5

1.1

1.0

0.7

4.5

0.33

0.25

13.52

0.09

Bone meal, raw

93.6

26.0

5.0

1.0

2.5

59.1

23.0510.22

4.16

—

Bone meal, raw, solvent process

93.1

25.7

1.0

1.9

63.5

24.0210.65

4.11

—

Bone meal, steamed

96.3

7.1

3.3

0.8

3.8

81.3

31.7415.00

1.14

0,18

Bone meal, steamed, solvent process

96.8

7.2

0.4

1.5

3.7

84.0

—

1.1b

—

Bone meal, steamed, special 97.7

Bone meal, 10 to 20% protein 97.2

Bread, white, enriched 64.1

Brewers’ grains, dried, 25% protein or over 92.9

Brewers’ qrains, dried, below 25% protein 92.3

13.5

14.6
8.5

27.6
23.4

7.9

6.5

2.0

6.5

6.4

1.0

1.5

0.3

14.3

16.1

5.1

3.6

52.0

40.9

42.5

70.2 31.8813.48 2.16
71.0 26.0012.66 2.34
1.3 0.06 0.10 1.36
3.6 0.29 0.48 4.42
3.9 - - 3.74

0.10

Brewers’ grains, dried, from California barley 91.1

Brewers’ grains, wet 23.7

Broom corn seed 89.7

Buckwheat, ordinary varieties 88.0

20.0

5.7

9.2

10.3

5.7
1.6

3.7
2.3

18.1

3.6

5.1

10.7

43.6

11.8

69.1

62.8

3.7

1.0

2.6

1.9

0.07

0.09

0.12

0.31

3.20

0.91

1.47

1.64

0.02

0.45

376/Appendix VI: A nalyses of Materials

Total dry

Protein

Fat

Fiber

N-free

Total

Calcium

Phos-

Nitro-

Potas-

Material

matter

extract i

minerals

phorus

gen

sium

Per ct.

Per c t.

Per ct.

Per rt.

Per ct.

Buckwheat, Tartary

88.1

10.1

2.4

12.7

60.9

2.0

0.13

0.31

1.62

0.44

Buckwheat feed, good grade

89.3

18.5

4.9

18.2

43.5

4.2

0.48

2.96

0.66

Buckwheat feed, low grade

88.3

13.3

3.4

28.6

39.8

3.2

—

0.37

2.13

0.68

Buckwheat flour

88.1

10.2

2.1

0.9

73.4

1.5

0.01

0.09

1.63

0.16

Buckwheat kernels, without hulls

88.0

14.1

3.4

1.8

66.5

2.2

0.05

0.45

2.26

0.49

Buckwheat middlings

88.7

29.7

7.3

7.4

39.4

4.9

—

1.02

4.76

0.98

Buttermilk

9.4

3.5

0.6

4.5

0.8

0.14

0.08

0.56

0.07

Buttermilk, condensed

29.7

10.9

2.2

12.6

4.0

0.44

0.26

1.74

0.23

Buttermilk, dried

92.4

32.4

6.4

0.3

43.3

10.0

1.36

0.82

5.18

0.71

Carob bean and pods

87.8

5.5

2.6

8.7

68.5

2.5

—

0.88

—

Carob bean pods

89.5

4.7

2.5

8.7

70.9

2.7

—

0.75

—

Carob bean seeds

88.5

16.7

2.6

7.6

58.4

3.2

—

2.67

—

Cassava roots, dried

94.4

2.8

0.5

5.0

84.1

2.0

—

0.45

—

Cassava meal (starch waste)

86.8

0.9

0.7

4.6

78.8

1.8

—

0.03

0.14

0.23

Cheese rind, or cheese meal

91.0

59.5

8.9

0.4

10.7

11.5

—

9.52

—

Chess, or cheat, seed

89.6

9.7

1.7

8.2

66.4

3.6

—

1.56

—

Chick peas

90.0

20.3

4.3

8.5

54.0

2.9

—

3.24

—

Citrus pulp, dried

90.1

5.9

3.1

1 1.5

62.7

6.9

2.07

0.15

0.94

—

Citrus pulp and molasses, dried

92.0

5.3

2.8

9.3

66.6

8.0

—

0.84

—

Citrus pulp, wet

18.3

1.2

0.6

2.3

12.8

1.4

—

0.19

—

Clover seed, red

87.5

32.6

7.8

9.2

31.2

6.7

—

5.22

—

Clover seed screenings, red

90.5

28.2

5.9

10.2

40.3

5.9

—

4.51

—

Clover seed screenings, sweet

90.1

21.7

3.7

14.7

41.1

8.9

—

3.47

—

Cocoa meal

96.0

24.3

17.1

5.1

43.7

5.8

—

3.89

—

Cocoa shells

95.1

15.4

3.0

16.5

49.9

10.3

—

0.59

2.46

2.16

Coconut oil meal, hydr. or exp. process

93.2

21.3

6.7

10.7

48.3

6.2

0.21

0.64

3.41

1.95

Coconut oil meal, high in fat

93.7

21.0

10.6

1 1.3

44.4

6.4

—

3.36

—

Coconut oil meal, solvent process

91.1

21.4

2.4

13.3

47.4

6.6

—

3.42

—

Cod-liver oil meal

92.5

50.4

28.9

0.7

9.6

2.9

0.18

0.61

8.06

—

Corn, dent, Grade No. 1

87.0

8.8

4.0

2.1

70.9

1.2

0.02

0.28

1.41

0.28

Corn, dent, Grade No. 2

85.0

8.6

3.9

2.0

69.3

1.2

0.02

0.27

1.38

0.27

Corn, dent, Grade No. 3

83.5

8.4

3.8

2.0

68.1

1.2

0.02

0.27

1.34

0.27

Corn, dent, Grade No. 4

81.1

8.2

3.7

1.9

66.2

1.1

0.02

0.26

1.31

0.26

Corn, dent, Grade No. 5

78.5

7.9

3.6

1.9

64.0

1.1

0.02

0.25

1.26

0.25

Corn, dent, soft or immature

70.0

7.2

2.3

2.5

56.5

1.5

—

0.24

1.16

0.26

Corn, flint

88.5

9.8

4.3

1.9

71.0

1.5

>— ■

0.33

1.57

0.32

Corn, pop

90.0

1 1.5

5.0

1.9

70.1

1.5

—

0.29

1.84

—

Corn ears, including kernels and cobs

(corn-and-cob meal)

86.1

7.3

3.2

8.0

66.3

1.3

—

0.22

1.17

0.29

Corn ears, soft or immature

64.3

5.8

1.9

7.8

47.7

1.1

—

0.93

—

Corn, snapped, or ear-corn chops with husks

88.8

8.0

3.0

10.6

64.8

2.4

“

—

1.28

—

Corn, snapped, very soft or immature

60.0

5.3

1.8

8.2

42.7

2.0

—

0.85

—

Corn bran

90.6

9.7

7.3

9.2

62.0

2.4

0.03

0.27

1.56

0.56

Corn feed meal

88.6

9.8

4.7

2.9

69.2

2.0

0.03

0.34

1.57

0.28

Corn germ meal

93.0

19.8

7.8

8.9

53.2

3.3

—

0.58

3.17

0.21

Corn gluten feed, all analyses

90.9

25.5

2.7

7.6

48.8

6.3

0.48

0.82

4.08

0.54

Appendix VI: Analyses of Materials/377

Material

Total dry Protein
matter

Per ct- Per ct.

Fat

Per ct.

Fiber
Per ct.

N-free
extract
Per ct.

Total
minerals
Per ct.

Calcium
Per ct.

Phos-
phorus
Per ct.

Nitro-
gen
Per ct.

Potas-
sium
Per ct.

Corn gluten feed, 25% protein guarantee

91.1

26.6

3.0

7.2

48.2

6.1

—

4.26

— „

Corn gluten feed, 23% protein guarantee

91.4

24.8

2.6

7.8

49.8

6.4

—

3.97

— *

Corn gluten feed with molasses

88.8

22.6

2.1

6.8

50.9

6.4

—

3.62

—

Corn gluten meal, all analyses

91.4

43.1

2.0

4.0

39.8

2.5

0.13

0.38

6.90

0.02

Corn gluten meal, 41% protein guarantee

91.4

42.9

2.0

3.9

40.1

2.5

—

6.86

—

Corn grits

88.4

8.5

0.5

0.6

78.4

0.4

—

1.36

—

Corn meal, degerminated, yellow

88.7

8.7

1.2

0.6

77.1

1.1

0.01

0.14

1.39

—

Corn meal, degerminated, white

88.4

8.6

1.2

0.7

76.1

1.8

0.01

0.14

1.38

—

Corn oil meal, old process

91.7

22.3

7.8

10.3

49.0

2.3

0.06

0.56

3.57

—

Corn oil meal, solvent process

91.7

23.0

1.5

10.4

54.6

2.2

0.03

0.50

3.68

—

Corn-starch

88.6

1 1 .6

0.6

0.1

0.2

87.6

0.1

—

0.10

Corn-and-oat feed, good grade

89.6

11.9

4.0

5.4

65.9

2.4

0.05

0.30

1.90

0.34

Corn-and-oat feed, low grade

89.6

9.1

2.9

13.4

59.0

5.2

—

1.46

—

Cottonseed, whole

92.7

23.1

22.9

16.9

26.3

3.5

0.14

0.70

3.70

1.11

Cottonseed, immature, dried

93.2

20.5

15.9

24.1

29.0

3.7

—

3.28

—

Cottonseed, whole pressed, 28% protein
guarantee

93.5

28.2

5.8

22.6

32.2

4.7

4.51

Cottonseed, whole pressed, below 28% protein

93.5

26.9

6.5

24.7

30.8

4.6

0.17

0.64

4.30

1.25

Cottonseed feed, below 36% protein

92.4

34.6

6.3

14.1

31.5

5.9

0.26

0.83

5.54

1.22

Cottonseed flour

94.4

57.0

7.2

2.1

21.6

6.5

—

9.12

Cottonseed kernels, without hulls

93.6

38.4

33.3

2.3

15.1

4.5

—

6.14

—

Cottonseed meal, 45% protein and over

93.5

46.2

7.7

8.6

24.9

6.1

0.22

1.13

7.39

Cottonseed meal, 43% protein grade, not
including Texas analyses

92.7

43.9

7.1

9.0

26.3

6.4

0.23

1.12

7.02

1.45

Cottonseed meal, 43% protein grade, Texas
analyses

92.5

42.7

6.4

10.6

27.0

5.8

0.19

0.96

6.83

1.34

Cottonseed meal, 41% protein grade, not
including Texas analyses

92.8

41.5

6.3

10.4

28.1

6,5

0.20

1.22

6.64

1.48

Cottonseed meal, 41% protein grade, Texas
analyses

92.1

41.0

6.0

1 1.6

27.6

5.9

—

— ,

6.56

—

Cottonseed meal, below 41% protein grade

92.4

38.2

6.2

12.3

29.4

6.3

0.23

1.29

6.1 1

1.57

Cottonseed meal, solvent process

90.8

44.4

2.6

12.7

24.3

6.8

—

7.10

—

Cowpea seed

89.0

23.4

1.4

4.0

56.7

3.5

0.1 1

0.46

3.74

1.30

Crab meal

92.4

31.5

2.0

10.7

5.0

43.2

15.15

1.63

5.04

0.45

Darso grain

90.0

10.1

3.1

1.9

73.5

1.4

0.02

0.32

1.62

—

Distillers’ dried corn grains, without solubles

92.9

28.3

8.8

11.4

41 .9

2.5

0.11

0.47

4.53

0.24

Distillers’ dried corn grains, with solubles

93.1

28.8

8.9

9.0

41.7

4.7

0.16

0.74

4.61

—

Distillers’ dried corn grains, solvent extracted

93.7

33.4

1.4

8.6

46.4

3.9

—

5.34

Distillers’ dried rye grains

93.9

18.5

6.4

15.6

51.0

2.4

0.13

0.43

2.96

0.04

Distillers’ rye grains, wet

22.4

4.4

1.5

2.5

13.3

0.7

—

0.70

—

Distillers’ dried wheat grains

93.7

28.7

6.1

13.0

42.2

3.7

—

4.59

—

Distillers’ dried wheat grains, high protein

94.7

46.2

5.7

10.9

30.0

1.9

—

7.39

—

Distillers’ solubles, dried, corn

93.0

26.7

7.9

2.6

48.4

7.4

0.30

1.41

4.27

1.75

Distillers’ solubles, dried, wheat

94.0

28.2

1.5

2.8

58.9

2.6

—

4.51

—

Distillery stillage, corn, whole

7.9

2.3

0.6

0.7

4.0

0.3 0.006 0.05

0.37

—

Distillery stillage, rye, whole

5.9

1.9

0.3

0.5

2.9

0.3

—

0.30

—

378/Appendix VI: Analyses of Materials

— — — — -

Total dry Protein

Fat

Fiber

N-free

Total

Calcium

Phos-

Nitro-

Potas-

Material

matter

extract

minerals

phorus

gen

slum

Per ct.

Distillery stillage, strained

3.8

1.1

0.4

0.2

1.8

0.3 0.004 0.05

0.18

Durra grain

89.8

10.3

3.5

1.6

72.4

2.0

—

1.64

—

Emmer grain

91.1

12.1

1.9

9.8

63.6

3.7

—

0.33

1.94

0.47

Feterita grain

89.4

12.2

3.2

2.2

70.1

1.7

0.02

0.33

1.96

Feterita head chops

89.6

10.7

2.6

7.4

65.7

3.2

—

1.71

Fish-liver oil meal

92.8

62.8

17.3

1.2

5.4

6.1

—

10.04

—

Fish meal, all analyses

92.9

63.9

6.8

0.6

4.0

17.6

4.14

2.67

10.22 0.40

Fish meal, over 63% protein

92.7

66.8

5.3

0.5

4.5

15.6

—

10.69

—

Fish meal, 58-63% protein

93.1

60.9

8.1

0.8

3.5

19.8

—

9.74

—

Fish meal, below 58% protein

93.2

56.2

11.0

0.7

2.9

22.4

—

8.99

—

Fish meal, herring

93.5

72.5

7.3

0.7

1.5

1 1.5

2.97

2.08

11.60

—

Fish meal, menhaden

93.6

62.2

8.5

0.7

4.2

18.0

5.30

3.38

9.96

—

Fish meal, redfish

94.2

56.7

11.4

0.9

24.3

4.01

2.44

9.07

—

Fish meal, salmon

92.8

59.4

9.8

0.3

4.3

19.0

5.49

3.65

9.50

—

Fish meal, sardine

93.1

67.2

5.0

0.6

5.4

14.9

4.21

2.54 10.76 0.33

Fish meal, tuna

90.1

58.2

7.9

0.7

3.4

19.9

4.80

3.10

9.31

—

Fish meal, whitefish

90.4

63.0

6.7

0.1

20.5

—

10.08

—

Fish solubles, condensed

49.5

29.3

8.4

—

2.2

9.6

—

4.69

—

Flaxseed

93.8

24.0

35.9

6.3

24.0

3.6

0.26

0.55

3.84

0.59

Flaxseed screenings

91.1

16.4

9.4

12.7

45.8

6.8

0.37

0.43

2.62

—

Flaxseed screenings oil feed

91.9

25.0

7.1

11.7

40.3

7.8

—

4.00

—

Garbage

39.3

6.0

7.2

1.1

22.2

2.8

—

0.96

—

Garbage, processed, high in fat

95.9

17.5

23.7

20.0

21.8

12.9

—

0.33

2.80

0.62

Garbage, processed, low in fat

92.3

23.1

3.5

13.5

38.1

14.1

—

3.70

—

Grapefruit pulp, dried

91.7

4.9

1.1

11.9

69.6

4.2

—

0.78

— -

Grape pomace, dried

91.0

12.2

6.9

30.2

36.7

5.0

—

1.96

Hegari grain

89.7

9.6

2.6

2.0

73.9

1.6

0.18

0.30

1.54

—

Hegari head chops

89.6

10.0

2.1

11.9

60.6

5.0

—

1.60

—

Hempseed oil meal

92.0

31.0

6.2

23.8

22.0

9.0

0.25

0.43

4.96

Hominy feed, 5% fat or more

90.4

11.2

6.9

5.2

64.2

2.9

0.22

0.71

1.79

0.61

Hominy feed, low in fat

89.7

10.6

4.3

5.0

67.4

2.4

—

1.70

—

Horse beans

87.5

25.7

1.4

8.2

48.8

3.4

0.13

0.54

4.1 1

1,16

Ivory nut meal, vegetable

89.4

4.7

0.9

7.2

75.5

1.1

—

0.76

—

Jack beans

89.3

24.7

3.2

8.2

50.4

2.8

—

3.96

—

Kafir grain

89.8

10.9

2.9

1.7

72.7

1.6

0.02

0.31

1.74

0.34

Kafir head chops

89.2

10.0

2.6

6.9

66.4

3.3

0.08

0.27

1.60

—

Kalo sorghum grain

89.2

11.8

3.2

1.6

70.9

1.7

—

1.89

—

Kaoliang grain

89.9

10.5

4.1

1.6

71.8

1.9

—

1.68

—

Kelp, dried

91.3

6.5

0.5

6.5

42.6

35.2

2.48

0.28

1.04

—

Lamb’s-quarters seed

90.0

20.6

4.5

15.1

40.2

9.6

—

3.30

—

Lespedeza seed, annual

91.7

36.6

7.6

9.6

32.8

5.1

—

5.86

—

Lespedeza seed, sericea

92.3

33.5

4.2

13.5

37.3

3.8

—

5.36

—

Lemon pulp, dried

92.8

6.4

1.2

15.0

65.2

5.0

—

1.02

—

Linseed meal, old process, all analyses

91.0

35.4

5.8

8.2

36.0

5.6

0.39

0.87

5.66

1.24

Linseed meal, o.p., 37% protein or more

90.9

38.0

5.9

7.7

33.7

5.6

0.39

0.86

6.08

1.10

Linseed meal, o.p., 33-37% protein

91.0

35.0

5.7

8.3

36.4

5.6

0.41

0.86

subset:

5.60

1.14

Appendix VI: Analyses of Materials/379

Total dry Protein

Fat

Fiber

N-free

Total <

Calcium

Phos-

Nitro-

Potas-

matter

extract r

minerals

phorus

gen

sium

Per ct.

Linseed meal, o.p., 31-33% protein

91.0

32.4

5.9

8.3

38.7

5.7

0.36

0.90

5.18

1.40

Linseed meal, solvent process, older analyses

90.4

36.9

2.9

8.7

36.3

5.6

—

5.90

—

Linseed meal and screenings oil feed

(linseed feed)

90.5

31.2

5.4

10.1

37.0

6.8

0.43

0.65

4.99

—

Liver meal, animal

92.3

66.2

16.4

1.4

1.9

6.4

0.62

1.27

10.59

—

Locust beans and pods, honey

88.4

9.3

2.4

16.1

57.1

3.5

—

1.49

—

Lupine seed, sweet, yellow

88.9

39.8

4.9

14.0

25.7

4.5

0.23

0.39

6.37

0.81

Malt, barley

90.6

14.3

1.6

1.8

70.6

2.3

0.08

0.47

2.29

—

Malt sprouts

92.6

26.8

1.3

14.2

44.3

6.0

—

4.29

—

Meat scraps, or dry-rendered tankage, 60%

protein grade

93.8

60.9

8.8

2.4

1.1

20.6

6.09

3.49

9.74

—

Meat scraps, or dry-rendered tankage, 55%

protein grade

93.9

55.8

9.3

2.1

1.3

25.4

8.33

4.04

8.93

—

Meat scraps, or dry-rendered tankage, 55%

protein grade, low fat

93.0

56.0

3.5

2.6

1.5

29.4

—

8.96

—

Meat scraps, or dry-rendered tankage, 52%

protein grade

93.1

52.9

7.3

2.2

4.3

26.4

—

8.46

T I ->

Meat and bone scraps, or dry-rendered tankage

with bone, 50% protein grade

93.9

51.0

10.1

2.1

1.6

29.1

9.71

4.81

8.16

—

Meat and bone scraps, or dry-rendered tankage

with bone, 45% protein grade

94.5

46.3

12.0

2.0

2.3

31.9

1 1.21

4.88

7.41

—

Mesquite beans and pods

94.0

1 3.0

2.8

26.3

47.4

4.5

—

2.08

—

Milk, cow’s

12.8

3.5

3.7

4.9

0.7

0.12

0.09

0.56

0.14

Milk, ewe’s

19.2

6.5

6.9

4.9

0.9

0.21

0.12

1.04

0.19

Milk, goat’s

12.8

3.7

4.1

4.2

0.8

0.13

0.10

0.59

0.15

Milk, mare’s

9.4

2.0

1.1

5.9

0.4

0.08

0.05

0.32

0.08

Milk, sow’s

19.0

5.9

6.7

5.4

1.0

—

0.94

—

Milk albumin, or lactalbumin, commercial

92.0

49.5

0.9

1.0

12.8

27.8

—

7.92

—

Milk, whole, dried

96.8

24.8

26.2

0.2

40.2

5.4

—

3.97

—

Millet seed, foxtail varieties

89.1

12.1

4.1

8.6

60.7

3.6

—

0.20

1.94

0.31

Millet seed, hog, or proso

90.4

1 1.9

3.4

8.1

63.7

3.3

0.05

0.30

1.90

0.43

Millet seed, Japanese

89.8

10.6

4.9

14.6

54.7

5.0

—

0.44

1.70

0.33

Milo grain

89.4

11.3

2.9

2.2

71.3

1.7

0.03

0.30

1.81

0.36

Milo head chops

90.1

10.2

2.5

6.9

66.2

4.3

0.14

0.26

1.63

—

Molasses, beet

80.5

8,4

62.0

10.1

0.08

0.02

1.34

4.77

Molasses, beet, Steffen’s process

78.7

7.8

62.1

8.8

0.1 1

0.02

1.25

4.66

Molasses, cane, or blackstrap

74.0

2.9

62.1

9.0

0.74

0.08

0.4b

3.6/

Molasses, cane, high in sugar

79.7

1.3

74.9

3.5

—

0.21

—

Molasses, citrus

69.9

4.0

0.2

61.3

4.4

—

0.64

—

Molasses, corn sugar, or hydrol

80.5

0.2

77.8

2.5

—

0.03

—

Mustard seed, wild yellow

95.9

23.0

38.8

5.0

23.6

5.5

—

3.68

Oat clippings, or clipped-oat by-product

92.2

8.8

2.3

25.3

44.9

10.9

—

1.41

—

Oat kernels, without hulls (oat groats)

90.4

16.3

6.1

2.1

63.7

2.2

0.08

0.46

2.61

0.39

Oat meal, feeding, or rolled oats without hulls

90.8

16.0

5.5

2.7

64.2

2.4

0.07

0.46

2.5b

0.37

Oat middlings

91.4

15.9

5.2

3.3

64.6

2.4

0.08

0.45

2.54

0.5 /

92.4 5.6 1.8 27.9 50.8 6.3 0.13 0.16 0.90 0.60

Oat mill feed

380/Appendix VI: Analyses of Materials

Total dry Prolein

Fat

Fiber

N-free

Total

Calcium

Phos-

Nitro-

Pot as-

Material

matter

extract

minerals

phorus

gen

sium

Per ct.

Oat mill feed, poor grade

92.4

4.3

1.8

30.5

50.2

5.6

„

0.69

Oat mill feed, with molasses

92.4

5.5

1.4

24.1

55.0

6.4

—

0.88

—

Oats, not including Pacific Coast states

90.2

12.0

4.6

1 1.0

58.6

4.0

0.09

0.34

1.92

0.43

Oats, Pacific Coast states

91.2

9.0

5.4

1 1.0

62.1

3.7

—

1.44

—

Oats, hull-less

90.0

15.4

4.2

2.6

65.7

2.1

* —

2.46

Oats, light weight

91.3

12.3

4.7

15.4

54.4

4.5

—

1.97

—

Oats, wild

89.0

12.7

5.5

15.2

50.9

4.7

—

2.03

—

Olive pulp, dried, pits removed

95.1

14.0

27.4

19.3

31.0

3.4

—

2.24

—

Olive pulp, dried, with pits

92.0

5.9

15.6

36.5

31.5

2.5

—

0.94

—

Orange pulp, dried

87.9

7.7

1.5

8.0

67.3

3.4

—

1.23

—

Palm-kernel oil meal

91.4

19.2

6.7

1 1.9

49.7

3.9

—

0.69

3.07

0.42

Palm seed, Royal

86.5

6.1

8.3

22.8

43.8

5.5

—

0.98

Palmo middlings

94.1

16.1

9.7

6.7

56.3

5.3

—

— r

2.58

Pea feed, or pea meal

90.0

17.7

1.4

23.7

43.7

3.5

2.83

—

Pea hulls of seeds, or bran

91.5

4.8

0.4

48.5

34.3

3.5

—

0.77

—

Pea seed, field

90.7

23.4

1.2

6.1

57.0

3.0

0.17

0.51

3.74

1.03

Pea seed, field, cull

89.7

24.8

2.5

7.1

52.0

3.3

—

3.97

—

Pea seed, garden

89.2

25.3

1.7

5.7

53.6

2.9

0.08

0.40

4.04

0.90

Peanut kernels, without hulls

94.6

30.4

47.7

2.5

11.7

2.3

0.06

0.44

4.86

—

Peanut oil feed

94.5

37.8

9.6

14.3

26.2

6.6

—

6.04

—

Peanut oil feed, unhulled, or whole pressed

peanuts

93.1

35.0

9.2

22.5

21.4

5.0

—

5.60

—

Peanut oil meal, old process, all analyses

93.0

43.5

7.6

13.3

23.4

5.2

0.16

0.54

6.96

1.15

Peanut oil meal, o.p., 45% protein and over

93.4

45.2

7.4

12.1

23.7

5.0

—

7.23

—

Peanut oil meal, o.p., 43% protein grade

92.8

43.1

7.6

13.9

23.0

5.2

6.90

—

Peanut oil meal, o.p., 41% protein grade

93.8

41.8

7.8

12.7

25.9

5.6

—

6.69

—

Peanut oil meal, solvent process

91.6

51.5

1.4

5.7

27.2

5.8

— ' ■

—

8.24

—

Peanut skins

93.8

16.3

23.9

1 1.8

39.1

2.7

—

2.61

—

Peanut screenings

93.6

23.8

11.5

18.9

33.0

6.4

—

3.81

—

Peanuts, with hulls

94.1

24.9

36.2

17.5

12.6

2.9

—

0.33

3.98

0.53

Peri lia oil meal

91.9

38.4

8.4

20.9

16.0

8.2

0.56

0.47

6.14

—

Pigeon-grass seed

89.8

14.4

6.0

17.3

45.8

6.3

—

2.30

— *

Pigweed seed

90.0

16.8

6.2

15.9

47.8

3.3

—

2.69

—

Pineapple bran, or pulp, dried

85.3

4.0

1.9

19.4

57.2

2.8

0.20

0.10

0.64

—

Pineapple bran, or pulp, and molasses, dried

87.4

3.9

1.0

15.9

63.4

3.2

— ■

—

0.62

Poppy-seed oil meal

89.2

36.6

7.9

11.6

20.7

12.4

—

5.86

—

Potato meal, or dried potatoes

92.8

10.4

0.3

2.0

75.8

4.3

0.08

0.22

1.66

1.97

Potato pomace, dried

89.1

6.6

0.5

10.3

69.0

2.7

—

1.06

—

Pumpkin seed, not dried

55.0

17.6

20.6

10.8

4.1

1.9

—

2.82

—

Raisin pulp, dried

89.4

9.6

7.8

16.1

50.6

5.3

—

1.54

—

Raisins, cull

84.8

3.4

0.9

4.4

73.1

3.0

.■

—

0.54

—

Rape seed

90.5

20.4

43.6

6.6

15.7

4.2

—

3.26

—

Rape-seed oil meal

89.5

33.5

8.1

10.8

30.2

6.9

—

5.36

—

Rice, brewers’

88.3

7.5

0.6

78.8

0.8

0.04

0.10

1.20

—

Rice, brown

87.8

9.1

2.0

1.1

74.5

1.1

0.04

0.25

1.46

—

Rice, polished

87.8

7.4

0.4

i-Z.'TZZ'.

0.4

79.1

0.5

0.01

0.09

1.18

0.04

aBTE-.ms

Appendix Vi: Analyses of Materials/381

Total dry Protein

Fat

Fiber

N-free

Total Calcium

Phos-

Nitro-

Potas-

matter

extract minerals

phorus

gen

sium

Per ct. 1

Per ct.

Per ct. Per ct.

Per ct.

Rice bran

90.9

12.5

13.5

12.0

39.4

13.5

0.08

1.36

2.00

1.08

Rice grain, or rough rice

88.8

7.9

1.8

9.0

64.9

5.2

0.08

0.32

1.26

0.34

Rice polishings, or rice polish

89.8

12.8

13.2

2.8

51.4

9.6

0.04

1.10

2.04

1.17

Rubber seed oil meal

91.1

28.8

9.2

10.0

37.6

5.5

—

4.61

—

Rye grain

89.5

12.6

1.7

2.4

70.9

1.9

0.10

0.33

2.02

0.47

Rye feed

90.4

16.1

3.3

4.6

62.7

3.7

0.08

0.69

2.58

0.83

Rye flour

88.6

1 1.2

1.3

0.6

74.6

0.9

0.02

0.28 1 /279 0.46

Rye flour middlings

90.6

16.5

3.5

4.2

63.1

3.3

—

2.64

—

Rye middlings

90.2

16.6

3.4

5.2

61.2

3.8

—

0.44

2.66

0.63

Rye middlings and screenings

90.4

16.7

3.8

6.1

59.5

4.3

—

2.67

—

Safflower seed

93.1

16.3

29.8

26.6

17.5

2.9

—

2.61

—

Safflower seed oil meal, from hulled seed

91.0

38.0

6.8

21.0

17.0

8.2

—

6.08

—

Safflower-seed oil meal from unhulled seed

91.0

18.2

5.5

40.4

24.1

2.8

—

2.91

—

Sagrain sorghum grain

90.0

9.5

3.5

2.1

73.4

1.5

0.43

0.39

1.52

—

Screenings, grain, good grade

90.0

15.8

5.2

9.2

54.3

5.5

—

2.53

—

Screenings, grain, chaffy

91.5

14.3

4.4

18.3

46.1

8.4

—

2.29

—

Schrock sorghum grain

89.1

10.2

3.0

3.4

70.8

1.7

—

1 .63

—

Sesame oil meal

93.7

42.8

9.4

6.2

22.8

12.5

2.02

1.61

6.84

1.35

Sesbania seed

90.8

31.7

4.3

13.5

38.0

3.3

—

5.07

—

Shallu grain

89.8

13.4

3.7

1.9

68.9

1.9

—

2.14

—

Shallu head chops

90.5

12.7

3.5

9.2

61.9

3.2

—

2.03

—

Shark meal

91.2

74.5

2.7

0.5

13.5

3.48

1.92

12.69

—

Shrimp meal

89.7

46.7

2.8

11.1

1.3

27.8

—

7.47

—

Skimmilk, centrifugal

9.5

3.6

0.1

5.1

0.7

0.13

0.10

0.58

0.15

Skimmilk, gravity

10.1

3.6

0.8

5.0

0.7

0.13

0.10

0.58

0.15

Skimmilk, dried

94.2

34.7

1.2

0.2

50.3

7.8

1.30

1.03

5.56

1.46

Sorghum seed, sweet

89.2

9.5

3.3

2.0

72.8

1.6

0.02

0.28

1.52

0.37

Soybean seed

90.0

37.9

18.0

5.0

24.5

4.6

0.25

0.59

6.06

1.50

Soybean flour, medium in fat

92.9

47.9

6.7

2.4

29.9

6.0

—

7.66

—

Soybean flour, solvent extracted

91.5

48.5

0.8

2.6

33.0

6.6

—

7.76

—

Soybean mill feed, chiefly hulls

90.8

11.8

2j 7 in

34.0

38.1

4.2

—

1.89

—

Soybean oil meal, expeller or hydraulic

/ '

process, all analyses

90.0

5.7

29.6

6.0

0.29

0.66

7.09

1.77

Soybean oil meal, exp. or hydr. process,

y >J

44-45% protein guarantee

91.3

45>i

5.4

29.3

5.9

0.31

0.68

7.26

1.92

Soybean oil meal, exp. or hydr. process,

43% protein guarantee

91.2

44.6

5.3

5.8

29.4

6.1

0.30

U.b/

7.14

—

Soybean oil meal, exp. or hydr. process,

41% protein guarantee

90.9

44.2

5.3

5.7

29.7

6.0

0.26

0.59

7.07

—

Soybean oil meal, solvent process

90.6

46.1

1.0

5.9

31.8

5.8

0.30

0.66

7.38

1.92

Starfish meal

96.5

30.6

5.8

1.9

14.3

43.9

—

4.90

—

Sudan-grass seed

92.4

14.2

2.4

25.4

38.4

12.0

—

2.27

—

Sunflower seed

93.6

16.8

25.9

29.0

18.8

3.1

—

0.55

2.69

0.66

Sunflower seed, hulled

95.5

27.7

41.4

6.3

16.3

3.8

0.20

0.96

4.43

0.92

Sunflower-seed oil cake, from unhulled seed,

solvent process

89.2

19.6

1.1

35.9

27.0

5.6

—

3.14

—

382/Appendix VI: Analyses of Materials

Material

Total dry Protein
matter

Per ct. Per ct.

Fat

Per ct.

Fiber
Per ct.

N-free Total Calcium

extract minerals

Per ct. Per ct. Per ct.

Phos-
phorus
Per ct.

Nitro-
gen
Per ct.

Potas-
sium
Per ct.

Sunflower-seed oil cake, from hulled seed,

hydr. process

90.6

36.3

13.5

14.2

20.2

6.4

0.43

1.04

5.81

1.08

Sweet clover seed

92.2

37.4

4.2

11.3

35.8

5.b

—

5.98

—

Sweet potatoes, dried

90.3

4.9

0.9

3.3

77.1

4.1

0.21

0.18

0.78

—

Tankaae or meat meal, digester process,

60% protein grade

93.1

60.6

8.5

2.0

1.8

20.2

6.37

3.23

9.70

0.46

Tankage with bone, or meat and bone meal,
digester process, 50% protein grade

93.5

51.3

11.5

2.3

26.1

10.97 5.14

8.21

—

Tankage with bone, or meat and bone meal,
digester process, 40% protein grade

94.7

42.9

14.1

2.2

4.1

31.4

13.49 5.18

6.86

Tomato pomace, dried

94.6

22.9

15.0

30.2

23.4

3.1

—

3.66

—

Velvet bean seeds and pods (velvet bean feed)

90.0

18.1

4.4

13.0

50.3

4.2

0.24

0.38

2.90

1.20

Velvet beans, seeds only

90.0

23.4

5.7

6.4

51.5

3.0

—

3.74

Vetch seed

90.7

29.6

0.8

5.7

51.5

3.1

—

4.74

—

Whale meal

91 .8

78.5

6.7

3.1

3.5

0.56

0.57

12.56

—

Wheat, average of all types

89.5

13.2

1.9

2.6

69.9

1.9

0.04

0.39

2.11

0.42

Wheat, hard spring, chiefly northern plains
states

90.1

15.8

2.2

2.5

67.8

1.8

—

2.53

—

Wheat, hard winter, chiefly southern plains

states

89.4

13.5

1.8

2.8

69.2

2.1

*—

—

2.16

—

Wheat, soft winter, Miss, valley and eastward

89.2

10.2

1.9

2.1

73.2

1.8

—

1.63

Wheat, soft, Pacific Coast states

89.1

9.9

2.0

2.7

72.6

1.9

—

1.58

—

Wheat bran, all analyses

90.1

16.9

4.6

9.6

52.9

6.1

0.14

1.29

2.70

1.23

Wheat bran, chiefly hard spring wheat

91.1

17.9

4.9

10.1

52.2

6.1

0.13

1.35

2.86

—

Wheat bran, soft wheat

90.5

16.1

4.3

8.7

55.7

5.7

—

2.58

—

Wheat bran, winter wheat

89.9

15.5

4.2

8.9

55.1

6.2

—

2.48

— *■

Wheat bran and screenings, all analyses

90.0

16.8

4.5

9.6

53.0

6.1

0.14

1.21

2.69

Wheat brown shorts

88.7

16.9

4.2

7.1

56.0

4.5

—

2.70

—

Wheat brown shorts and screenings

88.7

17.0

4.1

7.0

56.0

4.6

—

2.72

—

Wheat flour, graham

88.1

12.5

1.9

1.8

70.4

1.5

0.04

0.36

2.00

0.46

Wheat flour, low grade

88.4

15.4

1.9

0.5

69.7

0.9

—

“

2.4b

—

Wheat flour, white

88.0

10.8

0.9

0.3

75.6

0.4

0.02

0.09

1.73

U.U5

Wheat flour middlings

89.2

18.3

4.2

3.8

59.8

3.1

0.09

0.71

2.93

0.89

Wheat flour middlings and screenings

89.6

18.2

4.5

5.2

57.8

3.9

0.14

0.68

2.91

—

Wheat germ meal, commercial

90.8

31.1

9.7

2.6

42.2

5.2

0.08

1.11

4.98

0.29

Wheat germ oil meal

89.1

30.4

4.9

2.6

46.4

4.8

—

4.86

Wheat gray shorts

88.9

17.9

4.2

5.7

56.9

4.2

0.13

0.84

2.86

Wheat gray shorts and screenings

88.6

17.6

4.0

5.8

57.0

4.2

—

2.82

Wheat mixed feed, all analyses

89.7

17.2

4.5

7.2

56.1

4.7

0.11

1.09

2.76

—

Wheat mixed feed, hard wheat

89.8

18.7

4.8

7.7

53.6

5.0

0.1 1

1.09

2.99

Wheat mixed feed and screenings

89.3

17.5

4.3

7.1

55.7

4.7

0.1 1

0.96

2.80

—

Wheat red dog

89.0

18.2

3.6

2.6

61.9

2.7

0.07

0.51

2.91

O.bU

Wheat red dog, low grade

89.2

17.9

4.8

4.9

57.9

3.7

—

2.86

—

Wheat screenings, good grade

90.4

13.9

4.7

9.0

58.2

4.6

0.44

0.39

2.22

89.6 18.1 4.8 6.5 55.8 4.4 0.09 0.93 2.90 1.04

Wheal standard middlings, all analyses

Appendix Vi: Analyses of Materials/383

Total dry Protein

Fat

Fiber

N-free

Total Calcium

Phos-

Nitro-

Potas-

matter

extract minerals

phorus

gen

sium

Per ct.

Wheat standard middlings and screenings,

all analyses

89.7

18.0

4.7

7.4

55.1

4.5

0.15

0.88

2.88

—

Wheat white shorts

89.7

16.1

3.1

2.9

65.0

2.6

—

2.58

—

Whey, from cheddar cheese

6.9

0.9

0.3

5.0

0.7

0.05

0.04

0.14

0.19

Whey, skimmed

6.6

0.9

0.03

5.0

0.7

—

0.14

—

Whey, condensed

57.3

8.8

0.6

42.0

5.9

—

1.41

—

Whey, dried

93.5

12.2

0.8

0.2

70.4

9.9

0.86

0.72

1.96

—

Whey solubles, dried

96.3

17.5

2.0

62.8

14.0

—

2.80

—

Yeast, brewers’, dried

93.8

49.3

1.0

3.7

31.9

7.9

0.13

1.56

7.89

—

Yeast, irradiated, dried

93.9

48.7

1.1

5.5

32.2

6.4

0.07

1.55

7.79

2.14

Yeast, dried, with added cereal

90.2

12.3

3.7

3.2

68.5

2.5

0.09

0.45

1.97

Yeast, molasses distillers’, dried

91.0

38.8

1.9

6.1

30.2

14.0

—

6.21

—

(These tables have been adapted from “U.S. -Canadian Tables of Feed Composition”; Publication 1684;
Committee on Animal Nutrition and National Committee on Animal Nutrition, Canada, National Academy
of Sciences— National Research Council, Washington, D.C. 1969)

384/Appendix VII: Resources for Mushroom Growers

Suppliers of Cultures of Edible
Mushrooms

American Type Culture Collection
12301 Parklawn Drive
Rockville, Maryland 20852

Pennsylvania State University
2111 Buckhouf Laboratory
University Park, Maryland 16802

General Supplies & Equipment

Fungi Perfecti
PO Box 7634
Olympia, Wa. 98507
Phone: 360-426-9292
Fax: 360-426-9377

Products include: micron filters; laminar flow
hoods; sterile and room supplies; pressure
cookers; steam boilers; cultures; sterile and
growing room instruments; pure cultures
and spawn. Write for free brochure or in-
clude $4.50 for our fully illustrated
catalogue. International orders welcomed.

Commercial Spawn Makers

Rainforest Mushroom Spawn
International Division
PO Box 1 793
Gibsons, B.C.

Canada VON 1V0

Spawn of these species available: Pleurotus
ostreatus; Pleurotus sajor-caju; Pleurotus
cornucopiae; Pleurotus eryngii; Lentinus
edodes; and Stropharia rugoso-annulata.

Appendix VII: Resources for Mushroom Growers/385

Publications

The Mushroom Journal (monthly)

The Mushroom Growers Association (MGA)

Agriculture House

Knightsbridge

London, SW1X 7NJ

(Available in large university libraries.)

The Mushroom News (monthly)

American Mushroom Institute (AMI)

Box 373

Kennett Square, Pa. 19348
(Available in large university libraries.)

Mushroom Growing Bulletins
Dr. Paul Wuest
Department of Botany
Pennsylvania State University
University Park, Pa. 16802

Mi/cologia

The New York Botanical Garden
Bronx, New York 10458
Mushroom Newsletter for the Tropics
(quarterly)

The International Mushroom Society for the
Tropics

c/o Department of Biology

The Chinese University of Hong Kong

Shatin, N.T. Hong Kong

Organizations

North American Mycological Association
(NAMA)

4245 Redinger Rd.

Portsmouth, Oh. 45662

The Mycological Society of America (MSA)

c/o Dr. Harry Thiers

Department of Biology

San Francisco State University

1 600 Holloway Ave.

San Francisco, Ca. 94132

MycoMedia
PO Box 2222
Olympia, Wa. 98507
(Sponsors yearly forays and educational
conferences.)

Puget Sound Mycological Society
2161 E. Hamlin
Seattle, Wa. 98112

386/Appendix VIII: Metric Conversion Tables

Cropping Yield Substrate Depths

Ibs./sq. ft.

kg./sq. m.

inches

centimeters

1.0

4.9

2.54

1.4

6.8

5.08

2.0

9.8

7.62

2.2

10.8

10.16

2.6

12.7

12.70

3.0

14.7

15.24

3.5

17.7

17.78

4.0

19.6

20.32

4.5

22.0

22.86

5.0

24.4

25.40

5.5

27.0

27.94

6.0

29.0

30.48

6.5

31.4

Inch = 2.54 centimeters

Length

Centimeter = 0.3937 inches

Foot = 0.3048 meters

Meter = 3.2808 feet

Yard = 0.9144 meters

Meter = 1 .0936 yards

Sq. in. = 6.4516 sq. cm.

Area

Sq. cm. = 0.1550 sq. in.

Sq. ft. = 0.0929 sq. m.

Sq. m. = 10.7639 sq. ft.

Sq. yd. = 0.8361 sq. m.

Sq. m. = 1 .1 960 sq. yd.

Cu. in. = 16.3872 cu. cm.

Volume

Cu. cm. = 0.0610 cu. in.

Cu. ft. = 0.0283 cu. m.

Cu. m. = 35.3145 cu. ft.

Cu. yd. = 0.7646 cu. m.

Cu. m. = 1.3079 cu. yd.

Appendix VIII: Metric Conversion Tables/387

Cu. in. = 0.01 64 liters
Cu. ft. = 28.31 62 liters
Cup = 0.2366 liters
Quart = 0.9463 liters
Gallon = 3.7853 liters
A liter of distilled water weighs 1

Capacity

Liter = 61 .0250 cu. in.
Liter = 0.0353 cu. ft.
Liter = 0.2642 gal.

Liter = 1.0567 qt.

Liter = 1000 ml.
grams or 1 kilogram.

Grain = 0.0648 grams
Ounce = 28.3495 grams
Pound = 454 grams
Pound = 0.4536 kilograms
Ton = 907.1848 kilograms

Weight

Gram = 1 5.4324 grains
Gram = 0.0353 ounces
Kilogram = 2.2046 pounds
Metric Ton = 2,204 pounds

Pressure

1 kg. per sq. cm. = 14.223 lb. per sq. in.

1 lb. per sq. in. = .0703 kg. per sq. cm.

1 kg. per sq. m. = .2048 lb. per sq. ft.

1 atmosphere = 1.0332 kg. per sq. cm. = 4.696 lb. per sq. in. = 1.0133 bars

104

122

131

140

149

158

167

176

Temperature

Fahrenheit to Centigrade
5/9 ( °F - 32) = °C.
Centigrade to Fahrenheit
9/5 ( °C) + 32 = °F.

Miscellaneous Data

50 lbs. rye grain = 1 25 cups (approximately)
Area of a circle = tt (r) 2 = 3.14 x r 2

388 /Appendix VIII: Metric Conversion Tables

Glossary/389

acute Pointed, sharp.

adnate (of the gills) Bluntly attached to the

stem.

adnexed (of the gills) Attached to the stem
in an ascending manner
agar A product derived from seaweed and
valued for its gelatinizing properties. Com-
monly used to solidify media in any type of
sterile tissue culture.

Agaricales The order of mushrooms which
includes all mushrooms with true gills,
amyloid The characteristic bluish reaction
the flesh or the spores of a mushroom exhibits
in Melzer’s iodine.

anastomosis (pi. anastomoses) The
crosswise fusion of hyphal systems to form a
network of mycelia.

angstrom 10"'° microns, 1 ten thousandth
of a micron.

annular Resembling an annulus, appearing
as a ring-like zone.

annulus The tissue remnants of the partial
veil adhering to the stem and forming a mem-
branous collar or ring,
apex The top or highest point,
apiculus a term (misused) for the hilar ap-
pendix, the nipple-like projection by which a
spore is attached to the strerigmata (“arms”) of
the basidium.

appendiculate Hanging with fragments of
tissue.

appressed Flattened.

Ascomycetes Fungi that produce spores in
an ascus.

ascus A sac-like organ in which eight spores
develop and is characteristic of the perfect
stage of ascomycetous fungi,
autoclave A steam pressurized vessel used
to sterilize media.

bacillus (pi. bacilli) A general term for any
rod-shaped bacteria forming spores in free ox-
ygen environments. (The genus Bacillus is
more narrowly defined),
bacterium (pi. bacteria) The simplest
group of non-chlorophyll plant-like organisms.
Basidiomycetes All fungi which bear
spores externally upon a club-like cell known
as a basidium.

basidium, basidia A unique fertile cell,
club-like in form, in which meiosis occurs and
by which sexual spores are produced,
basidiocarp The fruiting body of fungi that
reproduce through basidia.
binucleate Having two nuclei in one cell,
border break The early occurence of
mushrooms along the edge of a substrate con-
tainer.

campanulate Bell shaped.

carpophore The fruiting body of higher

fungi.

390/Glossary

carpophoroid Analagous to a fruitbody,
although usually a poorly differentiated mass of
tissue, not well developed (“aborted”) and
sterile.

cartilaginous Brittle, not pliant,
casing A layer of water retentive materials
applied to a substrate to encourage and en-
hance fruitbody production,
caulocystidia Sterile cells covering the
mushroom stem.

cellular Composed of globose to rounded
cells, not thread-like.

cespitose Growing clustered, appearing to
arise from a single base,
cheilocystidia Sterile cells on the mush-
room gill edge, sometimes called marginal
cystidia.

chlamydospores A thick walled spore, typ-
ically of a secondary type, that arises directly
from the mycelium and having the full comple-
ment of chromosomes for producing off-
spring. Chlamydospores are typically ovoid
and heat resistant.

chrysocystidia A type of cystidia that is
highly refractive in once dried tissue revived
with a potassium hydroxide (KOH) solution
and appearing as a yellowish brown amor-
phous mass within the cell.

clamp connection An elbow-like protuber-
ance which arches over the walls separating
cells in mated (dikaryotic) mycelia of some
mushroom species.

compost A biological matrix of microorga-
nisms combined with straw, manure and other
organic substances and designed for mush-
room fruitbody production.

concolorous Having the same color.

conditioning The final conversion of
mushroom compost by selected microbial
groups.

conic Shaped like a cone.

conidia A uninucleate exteriorly borne cell

formed by constriction of the conidiophore.

conidiophore A specialized stalk arising
from mycelium upon which conidia are borne.

conidium (pi. conidia) An asexual spore
formed by the constriction of hyphae to chains
of cells.

context The flesh of a mushroom,
convex Regularly rounded.

Coprinaceae A family of mushrooms con-
taining the genera Coprinus, Panaeolus and
Psathprella.

coprophilous Growing on dung,
cortinate A type of veil consisting of fine
cobweb-like threads of tissue, extending from
the mushroom cap margin to the stem,
cropping The time of mushroom forma-
tion, development and harvesting.

cuticle The surface of cells on the cap that
can undergo varying degrees of differentiation.

cystidia Microscopic sterile cells adorning
the mushroom fruitbody.

decurrent The attachment of the gill plates
to the stem of mushrooms where the gills are
markedly downcurved, partially extending
down the stem.

decurved Curving in a downwards fashion,
deciduous Describing trees that seasonally
shed their leaves.

Glossary/391

deliquescing The process of autodigestion
by which the gills and cap of a mushroom melt
into a liquid. Most typical of the genus
Coprinus.

dikaryophase The phase in which there are
two individual nuclei in each cell of the mush-
room plant.

dikaryotic The state of cells in the
dikaryophase.

diploid A genetic condition where each cell

has a full set of chromosomes necessary for

sexual reporduction (2N).

disc The central portion of the mushroom

cap.

eccentric Off-centered,
ellipsoid Shaped like an oblong circle,
endospores Spores formed internally,
entheogenic A term to describe substances
that induce god inspiring feelings or expe-
riences.

equal Evenly proportioned,
eroded Irregularly broken,
evanescent Fragile and soon disappearing.

fibrillose Having fibrils,
fibrils Fine delicate ‘hairs’ found on the sur-
face of the cap or stem,
fibrous Composed of tough, stringy tissue,
filamentous Composed of hyphae or
thread-like cells.

flexuose, flexuous Bent alternately in op-
posite directions.

floccose Wooly tufts or cottony veil rem-
nants, typically adorning the cap or stem of
some mushroom species,
flush The collective formation and develop-
ment of mushrooms within a short time period,
often occurring in a rhythmic manner,
fructification The act of fruitbody forma-
tion.

fruitbody What is commonly called the
mushroom. The sexual reproductive body of
the mushroom plant.

fugacious Impermanent, easily torn or de-
stroyed.

fusoid Rounded and tapering from the cen-
ter.

gelatinous Having the consistency of jelly.
geno.ttfsfe The genetic heritage or constitu-
tlo^afpi organism. The genotype produces
fotype.

geotropism Growing oriented towards or in
response to gravity,
glabrescent Becoming smooth,
glabrous Smooth, bald,
glutinous Having a highly viscous gelati-
nous layer, an extreme condition of viscidity.
Gram (Gram’s Stain) A method for sepa-
rating bacteria whereby bacteria are stained
first with crystal violet (a red dye) and then
washed with an iodine solution. Gram positive
bacteria (Bacillus) retain the dye. Gram nega-
tive bacteria (Pseudomonas and some
Bacillus) lose the dye.

392/ Glossary

gregarious Growing numerously in small
groups but not in clusters.

habitat The substrate in which mushrooms
grow.

heliotropic Growing or turning towards the
sun.

heteromorphic Composed of different cell
types, usually describing the type of mush-
room gill edge.

heterothallic Having two or more morpho-
logically similar pairs of strains within the same
species. The combination of compatible spore
types is essential for producing fertile off-
spring. Typically a spore on a four spored ba-
sidium is compatible with only one of its coun-
terparts.

hilar appendix The stub-like protrusion on
the spore that connects the spore-producing
cell (ex. the basidium) to it.
hilum A marking on the spore where it was at-
tached to a spore-producing cell,
homomorphic Composed of similar cell
types.

homothallic Having one strain type that is,
by nature, dikaryotic and self-fertile; often aris-
ing from two spored basidia.

humicolous Growing in humus, soil,
hygrophanous Markedly fading in color
upon drying, used to describe the condition of
the mushroom cap.

hymenium The layer of fertile spore-
bearing cells on the gill.

hypha, hyphae Individual cells of
mycelium.

hyphal aggregate A concentration of
mycelium; a “knot” in the mycelial network
which often differentiates into a primordium.
hyphosphere The region immediately on
and surrounding hyphae.

indicator mold A mold, usually non-
destructive, whose occurrence indicates an im-
properly balanced condition in the substrate or
environment.

instar An insect in any of its stages of post
embryonic growth.

karyogamy The fusion of two sexually op-
posite nuclei within a single cell.

KOH Potassium Hydroxide, an agent com-
monly used to revive dried mushroom material
for microscopic study at a concentration of
2/2%.

lamellae Mushroom gills.

lignicolous Growing in wood or on a
substratum composed of woody tissue,
lignin The organic substance which, with
cellulose, forms the basis of most woody tis-
sue.

linear Considerably longer than wide, with
edges parallel,
lubricous Smooth

Glossary/393

lumen The amount of the flow of light emit-
ted from a single international foot candle,
lux The amount of illumination received by
a surface one meter from 1 foot candle, equal
to 1 lumen/square meter.

macroscopic Visible to the unaided eye.
matting A condition of a mycelium casing-
run that has become appressed from overwa-
tering. Similar to overlay except that matting
infers the mycelium has formed a dense, dead
layer of cells on the casing’s surface,
meiosis The process of reduction division
by which a single cell with a diploid nucleus
subdivides into four cells with one haploid
nucleus apiece.

membranous Being sheath-like in form,
mesophile An organism thriving in moder-
ate temperature zones, usually 40-90 °F.
metuloid Used to describe a sterile cell en-
crusted with a crystalline (calcitized) substance,
micron One millionth of a meter, 10' 6
meters, one thousandth of a millimeter.

microscopic Visible only with the aid of a
microscope.

mitosis The non-sexual process of nuclear
division in a cell by which the chromosomes of
one nucleus are replicated and divided equally
into two daughter nuclei,
monkaryon, monocaryon (adj. monokar-
yotic, monocaryotic) The haploid state of
the mushroom mycelium, typically containing
a single nucleus.

mottled Spotted, as in the uneven ripening
of spores on the gill surfaces that so character-
izes species in the genus Panaeolus.

mushroom A fleshy fungus that erects a
body of tissue in which sexual spores are pro-
duced and from which they are distributed.

mycelium (pi. mycelia) A network of
hyphae.

mycology The study of fungi,
mycophagist A person or animal that eats
fungi.

mycophile a person who likes mushrooms,
mycophobe a person who fears mush-
rooms.

mycorrhizal A peculiar type of symbiotic
relationship a mushroom mycelium forms with
the roots of a seed plant, typically trees.

nanometer 10" meters, one thousandth of
a micron.

natural culture The in vitro cultivation of
mushrooms by transplanting living mycelium,
usually from a natural habitat,
nomenclature Any system of classification,
nucleate Having nuclei
nucleotide One of the four nitrogenous
bases in DNA; often called the building blocks
of the DNA molecule.

nucleus, nuclei A concentrated mass of dif-
ferentiated protoplasm in cells containing chro-
mosomes and playing an integral role in the
reproduction and continuation of genetic mate-
rial.

obtuse Bluntly shaped.

394/Glossary

ochraceous Light orangish brown to pale
yellowish brown.

oidia Conidia (spores) borne in chains,
olivaceous Olive gray-brown,
overlay A condition of the casing where my-
celium been allowed to completely cover the
surface. Overlay is caused by prolonged vege-
tative growth temperatures, high C0 2 levels
and excessive humidity. Overlay, if overwa-
tered, becomes matted,
ovoid Oval shaped.

pallid Pale in color.

parasite An organism living on another liv-
ing species and deriving its sustenance to the
detriment of the host.

partial veil The inner veil of tissue extend-
ing from the cap margin to the stem and at first
covering the gills of mushrooms,
pasteurization A process by which bulk
materials are partially sterilized through contact
with live steam, hot water or dry heat at tem-
peratures of between 140-1 60 °F.

pellicle A skin-like covering on the cap,
sometimes gelatinous and separable.

pencillate Resembling a broom or brush;
Penicillium- like.

perithecium A flask shaped or pear shaped
saclike fruitbody of some Ascomycetes that en-
closes asci.

persistent Not disappearing with age.

Phase I The steps taken in the outdoor
preparation, assemblage and conversion of
raw materials into a nutritious medium for
mushroom growth.

Phase HI The pasteurization and final condi-
tioning of a mushroom compost,
phenotype The observable physical charac-
teristics resulting from interaction between the
host environment and the genotype,
photosensitive Sensitive to light,
phototropic Growing towards light,
pileus The mushroom cap.
pith The central cottony “stuffing” in the
stems of some mushrooms.

pleurocystidia Sterile cells on the surface
of mushroom gills. Sometimes called facial
cystidia.

pliant Flexible.

pore A circular depression at the end of
spores in many species.

primary mycelium The haploid and uninu-
cleate mycelium originating from the germina-
tion of a spore which is otherwise not capable
of producing a sporuiating organ.

primordium (pi. primordia) The first rec-
ognizable but undifferentiated mass of hyphae
that develops into a mushroom fruitbody. Syn-
onymous with “pinhead”.

pruinose Having a powdery appearance.

pseudorhiza A long root-like extension of
the stem.

pseudosclerotium (pi. pseudosclerotia)

A conglomerated mass of mycelial tissue re-
sembling a sclerotium but which can not pro-
duce a fruitbody or new mycelial growth.

psilocybian Having psilocybin and/or psi-
locin. (Not necessarily belonging to the genus
Psilocybe).

psilocyboid Resembling a Psilocybe mush-
room.

Glossary/395

radicate Tapering downwards. Having a
long root like extension of the stem,
reticulate Marked by lines or ridges,
rhizomorphs Cord-like or strand-like
hyphae.

SACing Nutrient supplementation of bulk
substrates at the time of casing,
saprophyte A plant (fungus) that lives on
dead organic matter.

saltation The mutation developing from an
isolate of mycelium having a known pure
genotype.

scanning electron microscope An elec-
tronic microscope which scans an object in a
vacuum with a beam of electrons resulting in
an image of high resolution and magnification
that is then displayed on a television-type mon-
itor.

sclerotium (pi. sclerotia) A hardened
mass of mycelium, usually darkly pigmented,
that is the resting (vegetative) phase in some
fleshy and non-fleshy fungi, and from which
fruitbodies or viable mycelium can arise,
scratching Ruffling of the substrate or cas-
ing surface in order to maintain an open, por-
ous condition conducive to primordia forma-
tion.

seceding Describing the condition where
the gills have separated in their attachment to
the mushroom stem and have torn free, usual-
ly leaving longitudinal ridges at the stem’s
apex.

sector A geometric growth of diverging my-
celium (most visible on media filled petri
dishes), the appearance of which contrasts with
that of neighboring mycelia, usually indicative
of genetic mutation.

secondary mycelium A dikaryotic and bi-
nucleate mycelium characterized by clamp
connections, crossing (anastomosis), and
which is assimilative, not generative, in func-
tion.

senescent Having grown old.
septate Having walls dividing cells.

sinuate Describing the attachment of the
mushroom gill to the stem at the junction of
which the gill appears notched.

somatic Being in the assimilative phase of
mycelial growth.

sordid Dirty looking,
spawn The aggregation of mycelium on a
carrier material which is usually used to inocu-
late prepared substrates,
spores The reproductive cells or “seeds” of
fungi, bacteria, and plants.

sporocarps Any truitbody that produces
spores.

sterigmata (pi. sterigmatae) Elongated ap-
pendages or “arms” extending from the apex
of the basidium and upon which spores form.

stipe The stem or stalk of a fungus.

strain A race of individuals within a species
sharing a common genetic heritage but differ-
ing in some observable features of no taxo-
nomic significance.

stroma A dense, cushion-like aggregation
of mycelium forming on the surface of com-
posts or casings and indicative of somatic
(vegetative), not generative growth.

396/Glossary

Strophariaceae The family of dark brown
spored mushrooms containing the genera
Melanotus, Naematoloma, Pholiota,
Psilocybe and Stropharia.
stropharioid Resembling a species of
Stropharia, i.e. having a membranous ring on
the stem, a convex cap, and purplish brown
spores.

substrate Straw, sawdust, compost, soil, or
any fibrous material on which mushrooms
grow.

terrestrial Growing on the ground.

tertiary mycelium Mycelium arising from
secondary mycelium that is involved in mush-
room fruitbody formation,
tetrapolar Having or located at four poles,
as with the four spored basidia in most mush-
rooms.

thermogenesis The process of heat gener-
ation by microorganisms,
thermophile An organism thriving in
75-1 40 °F. temperature zone.

trama The fleshy part of the cap between
the cap cuticle and the fertile spore bearing lay-
ers of the mushroom gill,
translucent Transmitting light diffusely,
semitransparent.

umbo A knob-like protrusion on the top
center of the mushroom cap.
umbonate Having an umbo,
uncinate A type of gill attachment,
undulating Wavy.

universal veil An outer layer of tissue envel-
oping the cap and stem of some mushrooms,
best seen in the youngest stages of fruitbody
development.

variety A sub-species epithet used to de-
scribe a consistently appearing variation of a
particular mushroom species,
vector The pathway by which a disease is
spread; the “vehicle” for distributing a patho-
gen.

veil A tissue covering mushrooms as they
develop.

vesicle A small bulb-like swelling (some-
times bladder-like).

viscid Slimy or slippery when moist, sticky
to the touch when partially dry. A characteristic
of the mushroom cap or stem.

zonate Having a band-like region darker in
color or different in form than the surrounding
tissue.

umbilicate Depressed in the center region
of the mushroom cap.

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Index/ 409

abnormalities, 152
abort, 147

Actinomyces, 96, 246
activated charcoal, 28
adenosine monophosphate, 358
aerobic, 84
algae, 355
agar media, 1 9, 20
Agaricus bitorquis, 1 6 1
Agaricus brunnescens, 140, 164
air, 66

air circulation, 71, 151
air exchanges, 69, 97, 152
alleles, 334
Alternaria, 257
ammonia, 82, 104
anaerobic, 84, 87
anastomosis, 8, 27
antibiotics, 20
Artbrobotrys, 332
Aspergillus, 259

Bacillus, 248

bacteria, 28-29, 43, 96, 248

bacterial blotch, 153, 252

bacterial pit, 252

bags (plastic), 116, 119

basidium, 4, 1 2

bifactorial mating system, 337

biomass, 97

bipolar, 1 67

Botrytis, 262

botulism (see Clostridium)
bran, 54

breeding (see Genetics)

brown mold (see Botrytis, Papulospora)

buffers, 131

bulk pasteurization, 101
bulk room, 1 02-1 03

calcium carbonate, 47, 81, 131
calcium sulfate, 47, 81
carbon dioxide, 126, 129, 143
caramellization, 21, 89
casing, 127
cecids, 325
cellulose, 78, 1 1 5
Chaetomium, 97, 264
chicken manure, 79-80
chlorine, 155
chromosome, 334
Chrysosporium, 226
Cladosporium, 268
Clostridium, 250
clamp connection, 8, IQ
cloning, 29
C:N ratio, 83

cobweb mold (see Dactylium)
compost, 77
turning, 87
conditioning, 97
cooldown, 104
contaminants, 233
Copelandia, 13, 183
Coprinus comatus, 1 68
Coprinus sp. 226, 270
cottony mycelium, 33, 35
cow manure, 80
cropping, 150
Cryptococcus, 273

Dactylium, 275

410 /Index

Damping off disease (see Fusarium)

delayed release nutrients, 126

dehydrator, 1 56

depth rings, 1 35

die back disease (see Virus)

dikaryon, 27

dikaryotization, 7

Doratomyces, 277

DNA, 334

dry bubble (see VerticiHium)
drying, 156
ducting, 70

endospore, 250
Epicoccum, 279
express compost, 106

fans, 68
filling, 98

filters, filtration, 18, 45, 70, 72
firefang (see Actinomyces)
Flammulina velutipes, 115, 172
flies, 97, 320
flushes, 1 50
formulas
agar, 20
air exchange, 69
spawn, 46
compost, 81-82
casing, 132
freeze drying, 1 56
fruitbody, 4
fruiting, 140

fungus gnats (see Sciarids)
Fusarium, 281

gametes, 335

genes, 333
genetics, 334
genotype, 334
Geotrichum, 284
germination, 8, 24
germ pore, 7
glove box, 17,19
grains, 42-43
Gram positive, 250
Gram negative, 355
greenhouse, 62
growing room, 61 , 98
gypsum (see Calcium sulfate)

haploid, 6
harvesting, 155
heating, 73
hemicellulose, 78
HEPA filters (see Micron)
heterothallic, 337
homothallic, 337
horse manure, 78, 81
Humicola, 96, 286
humidistats, 74
humidity, 74, 140, 144
hygrometer, 76
hyphae, 7

hyphal aggregate, 34
hyphosphere, 355
Hypomyces ( see Dactylium)

initiation (pinhead), 140-141
incubation, 57
inky caps (see Coprinus)
inoculation, 48, 51
instar, 321
insulation, 63
interferon, 345
illegitimate mating, 337

Index/41 1

karyogamy, 9
knotting, 143

laminar flow systems, 347

Lentinus edodes, 113-114, 176

Lepista nuda, 180

lignin, 78, 1 1 4

lignicolous fungi, 1 1 4

lights, 74, 1 47

lime (see Calcium carbonate)

liquid inoculation, 55-56

liquid nitrogen, 39

log culture, 114, 177

long composting, 89

natural culture, 1 1 0
nematodes, 97, 130, 331
neopeptone, 20
Neurospora, 296
nitrogen, 79, 82-83

olive green mold (see Chaetomiurri)
overlay, 143
oxygen, 68, 83

oyster mushroom (see Pleurotus)
oyster shell, 1 32

malt agar, 20

mating type, 336

media preparation, 1 9

medicinal properties, 343

meiosis, 335

mesophiles, 88

microclimate, 129

micron filters, 348

microorganisms, 79, 96, 129, 353

microporous filters, 45, 1 16

mini-logs, 116, 179

mites, 328

mitosis, 335

Monilia, 288

monokaryon, 27, 340

Mucor, 290

multispore culture, 24

mushroom extracts, 357

mushroom life cycle, 5-6

mutation, 334

Mycogone, 294

Mycelia Sterilia, 292

Myceliopthora, 267

mycelium, 4, 6, 1 0, 25

Panaeolus cyanescens, 127, 183

Panaeolus subbalteatus, 1 86

Papulospora, 298

pasteurization, 97, 130

patching, 142

pathogens, 235

peat moss, 131

Penicillium, 300, 348

perithecium, 263, 295

perlite, 54

pests, 31 8

pH, 20, 130

Phase I composting, 78

Phase II composting, 78, 96, 97, 100

phenotype, 334

phorids, 323

photosensitive, 1 47

phototropism, 147

picking, 155

pig manure, 80

pinheads (see primordia)

pin molds (see Rhizopus, Mucor)

Pleurotus ostreatus (type variety), 115, 189
Pleurotus ostreatus (Florida variety), 193
potatoe dextrose agar, 20

412 /Index

preservation, 37, 1 56
pressure cooker, 20
primary mycelium, 6
primorida, 4, 1 39
protein, 78, 82

Pseudomonas, 192, 251, 354-355
Psilocybe cubensis, 27, 59, 1 96
Psilocybe cyanescens, 1 1 0, 200
Psilocybe mexicana, 59, 204
Psilocybe tampanensis, 207

recombination, 335
Reishi, 345
Rhizobium, 355
rhizomorphic, 33
Rhizopus, 302
ricking, 85
ridge bed, 1 1 1
RNA, 333
rosecomb, 1 52

sacing, 1 26
sand, 1 30
sawdust, 54, 1 1 4
scaling, 151
Scenedesmus, 355
sciarids, 321

sclerotia, 205, 207, 297
Scopulariopsis, 304
scratching, 1 36
secondary mycelium, 6
sectoring, 31, 33, 338
senescence, 35
Sepedonium, 306
shelf growing, 64
shiitake (see Lentinus)
short composting, 90
Sinden. J.W., 42, 65-66, 90
single spore isolation, 340

slants, 37, 39
spawn, 42, 45, 54
spawning, 122
spawn running, 1 22
spore, 6, 24

spore dilution technique, 340
spore germination, 5, 24
spore printing, 23
static pressure, 68
sterigmata, 11-12
sterile room, 1 6
sterilization, 21 , 116
stock cultures, 37
Stoller, B.B., 33
strains, 31 , 36
strain generation, 35
straw, 78, 1 1 7
Streptomyces, 255
stroma, 35, 143

Stropharia rugoso-annulata, 26, 21 1
Stysanus (see Doratomyces)
substance X, 357
substrate preparation, 78, 1 10
sugar cane bagasse, 106
super spawning, 1 26
supplements, 79-81
supplementation
filling, 95
spawning, 126
casing, 1 26

synthetic composts, 79, 89, 91

temperature, 57, 59, 88, 122, 150

tertiary mycelium, 6

tetrapolar mating system, 337

thermogenesis, 84

thermometers, 76

thermophiles, 88, 96

thermostats, 74

tissue culture, 29

Torula, 96, 308

tray system, 65

Trichoderma, 3 1 0
Trichothecium, 3 1 3

unifactorial mating system, 337
uninucleate, 6
urea, 80

vector, 234

ventilation, 68, 72, 103
vermiculite, 132
Verticillium, 3 1 5
virgin spawn, 1 1 1
virus, 244

Volwariella volvacea, 2 1 4

watering, 85, 136, 154

wet bubble (see Mycogone)

wet spot (see Bacillus)

wheat straw, 78, 117

white plaster mold (see Scopulariopsis)

windrow, 84-85
wood substrates, 1 1

yeast, 28, 43

yellow mat disease (see Chrysosporium)
yield, 122, 150

414/Credits

Michael Beug
Fig. 157.

J.S. Chilton

Figs.: 45, 56, 64, 65, 67, 68, 70, 73-76, 82, 83, 84, 85, 86-91, 94, 95-100, 106, 107,
117, 119, 122, 126, 132, 133, 134, 143, 171, 237.

Anthoinette Gunter

Figs.: 1 , 2, 21 , 55, 1 97.

Rick Kerrigan

Figs.: 0, 108, 109, 151, 152, 153, 170.

Tom Lind with Calligraphy by Karen Porter.

Figs.: 77-80, 1 78, 1 79, 1 83, 1 86, 1 87, 1 89, 1 92-1 95, 1 98, 1 99, 201 , 202, 203, 205,
206, 207, 209, 211, 212, 214, 216, 217, 219, 221, 222, 224, 225, 227, 228, 231-
235, 238.

Mary Montoya

Figs.: 239, 241.

Chris Nelson

Figs.: 47, 69, 81, 163.

Cruz Stamets
Fig. 22.

Bill Wright
Fig 177.

Paul Stamets

3-20, 23-44, 46, 48-54. 57-62,66,71,72,92,93, 101-105, 1 1 1 -1 1 6, 1 1 8, 1 20, 1 21 ,
123-25, 127-131, 135-142, 144-150, 154-156, 158-162, 164-169, 173-176, ISO-
185, 188, 190, 191, 196, 200, 204, 208, 210, 213, 215, 218, 220, 223, 226, 229,
230, 236, 240, 242. Color plates 1-23 and cover photographs.

Acknowledgements/ 415

The authors are indebted to many who have made this work possible. Foremost amongst those
are our spouses Cruz Stamets and Janice Leighton. They provided more than just technical assis-
tance. Their encouragement, companionship and endurance has been critical to the success of this
project.

Besides the artists and photographers whose contributions are most appreciated, we would like
to express our thanks to John Stamets for editing the manuscript and printing the black and white
photographs, to Mike McCaw for his chapter on genetics, to Rick Kerrigan for his unwavering sup-
port, to Dr. Michael Beug for his encouragement and sponsorship, to The Evergreen State College
for the use of the scanning electron microscope, to Bill Street of Ostrom Mushroom Farms for his
generosity in allowing photographs to be taken of his facilities, to Kathy Harvey and Steve Raynor
for their contributions to the contamination section, to P.J.C. Vedder for his tutelage and contribu-
tion to practical mushroom cultivation, to Dr. Daniel Stuntz for his guidance and his teaching of tis-
sue culture techniques and to Chris W. Nelson of Sound Media Productions for his work in photo-
graphic development and printing.

Others who have helped us, in one way or the other, to complete this arduous endeavor are
Janet McCaw, Paul Kroeger, Dale Leslie, Heidi Stamets, Mike Maki, Jim Jacobs, William Raves,
Bob Chieger, David Tatelman, Samuel and Victoria, Mike, Michael, Gary, and mycophiles too
numerous to mention who, through their questions and curiosity, have determined much of the
direction, content and scope of this book. We thank you.

Hwa Rang

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